Unraveling the Spatio-Temporal Dynamics of Soil and Root-Associated Microbiome in Texas Olive Orchards: A Comprehensive Analysis

doi:10.21203/rs.3.rs-4066985/v1

Download PDF

Article

Unraveling the Spatio-Temporal Dynamics of Soil and Root-Associated Microbiome in Texas Olive Orchards: A Comprehensive Analysis

https://doi.org/10.21203/rs.3.rs-4066985/v1

This work is licensed under a CC BY 4.0 License

Journal Publication

published 06 Aug, 2024

Read the published version in Scientific Reports →

You are reading this latest preprint version

Understanding the structure and diversity of microbiomes is critical to establishing olives in non-traditional production areas. Limited studies have investigated soil and root-associated microbiota dynamics in olives across seasons or locations in the United States. We explored the composition and spatiotemporal patterns in two niches (roots and soils), seasons (spring, summer, and fall), and domains (bacteria and fungi) in the microbiome of the olive variety Arbequina across three olive orchards in Texas to investigate the structure of the olive-associated microbial communities and specificity to the root endosphere and soil rhizosphere zones. The bacterial populations in the rhizosphere (16.42%) and endosphere (15.49%) were dominated by Phylum Proteobacteria, followed by Actinobacteriota (RS, 12.63%; RE, 16.47%). Rubrobacter (5.27%) and Actinophytocola (3.49%) were dominant taxa in the rhizosphere and root endosphere at the genus level. Among fungal communities, phylum Ascomycota was prevalent in the rhizosphere (71.09%) and endosphere (41.37%). Members of the Chaetomiaceae taxon outnumbered (17.61%) another taxon in the root endosphere. As Per the alpha diversity indices, rhizosphere soil at Moulton showed much higher richness and diversity than other places, which predicted a significant difference in rhizosphere between locations for bacterial diversity and richness. There was no significant variation in the bacterial diversity in the niches and the fungal diversity within the root endosphere between locations. Beta diversity analysis confirmed the effect of compartments (Fungi: 12.3%; Bacteria: 45.1%) in influencing community differences. Microbial diversity was apparent within the endosphere (Bacteria:14.6%, Fungi:15.6%) and rhizosphere (30.5%, Fungi: 21.6%). The seasons influenced only the rhizosphere fungal diversity (8.5%), contrasting the bacterial diversity in either niche. The research provided a comprehensive overview of the microbial diversity present in both the rhizosphere and endosphere of olive trees. The abundance and composition of OTUs associated with the rhizosphere soil of Arbequina suggest its role as a source reservoir in defining the potential endophytes.

Biological sciences/Plant sciences

Biological sciences/Ecology/Agri ecology

Biological sciences/Ecology/Biodiversity

Olive (Olea europaea L.) is cultivated commercially for the quality of its oil worldwide. Although early domestication of olive trees began in the Mediterranean regions [1], in recent decades, commercial olive production expanded to non-traditional areas, such as Australia and North and South America [2], significantly varying in the agro-climatic conditions [3–5]. Olives were introduced into the United States in the late 18th century [6]. The United States only represents less than 5% of the global olive production [7]. While California is the primary center for olive cultivation, the industry spans other states, including Texas, Arizona, Georgia, Florida, Oregon, and Hawaii. In Texas, beginning in the mid-1990s [8], the high-density olive planting for oil production has been spread across approximately 1,400 ha [9]. The productivity of Texas olives is primarily affected by stress from cold or heat and diseases, prominently the cotton root rot (Phymatotricopsis omnivora), which is prevalent in the high-pH soils of southwest Texas. Even if olive trees are adapted to drought and maintain an ability to produce fruit in extreme climates [10], the changing climate conditions have shown varied impacts on olive fruit maturation and oil composition in different cultivars, locations, and water availability [11–13].

Soil and plant-associated microbial communities are critical to plant productivity. Plant species, selection pressure, and environmental conditions define diversity within such communities. Soil microbes regulate the mineralization and competition of nutrients that sustain plant productivity [14]. At the same time, plant-associated microbiomes confer fitness advantages to the plant host, including growth promotion, nutrient uptake, stress tolerance, and resistance to pathogens [15]. In return, plants can affect soil microbial communities via host preference and changes in plant-derived inputs, such as litter, rhizodeposits, and root exudates [16].

Several reviews have highlighted the significance of soil and olive tree-associated microbiomes in defining olive tree's productivity [17, 18]. However, since the world's production center for olives has been the Mediterranean basin, research on the interactions with soil microbiota and/or olive tree microbiomes outside these non-traditional production areas is missing. Microbiota, particularly at the root level, are critical in modifying plant physiology and metabolism under various climatic conditions [19]. The role of plant genotype in shaping the composition of its root-associated microbiome has been highlighted based on the differences and similarities between the microbial communities in different soils [20]. Healthy and highly stable root microbiota are critical in helping olive trees thrive in new environments and climatic conditions [21]. A study on the microbiomes (communities of bacteria and fungi) of the endo- and rhizosphere of various olive cultivars from the World Olive Germplasm Collection (WOGC) at the Institute of Agricultural and Fisheries Research and Training (IFAPA, Córdoba, Spain) revealed a robust genotypic influence and lower diversity in the endosphere than in the rhizosphere [20]. The changing dynamics of global climate (i.e., reduced rainfall, increasing drought and temperature) will likely expand the arid and semiarid environments characterized by low soil nutrients and organic matter, like the Texas climate, impacting olive productivity, soil microbiomes, and ecosystem functioning. A study evaluating the impact of aridity on the bulk soil and the olive root-associated bacterial communities indicated that with the increment of aridity, distinct bacterial communities dominated by aridity-winner and aridity-loser bacteria negatively and positively correlated with increasing annual rainfall [22]. Likewise, a study on the fungal communities in plant tissues and the influence of seasonality in olive trees suggested higher diversity in roots and spring than in summer [23]. The impact of variables such as plant age, altitudinal gradient, and geographic location, but the cultivar on the microbial communities of commercial olive plants has been demonstrated [24]. On the contrary, studies have also shown the effect of the olive cultivar on the distinct differences in the endophytic and epiphytic microbial communities [25, 26].

A deeper understanding of the structure and diversity of the soil and root microbiome in olive production would enable its utilization for abiotic or biotic stress alleviation, especially in non-traditional production areas. To our knowledge, no systematic investigation has been performed on this cultivar's soil/root-associated microbiota dynamics across seasons or locations in the United States or Texas. Here, we characterize the composition and spatiotemporal patterns in two niches (roots and soils), seasons (Spring, summer, and fall), and domains (bacteria and fungi) in the microbiome of olive tree variety 'Arbequina' the primary variety used for commercial production in the United States planted at three locations (Carrizo Springs, Moulton, and Berclair) across Texas. We hypothesized that (a) the sampling location and seasons will strongly structure the olive-associated microbial communities over broadscale changes in climate and soil features across Texas, (b) the root microbial community (endosphere) structure would be more responsive than soil communities over the seasons or locations and (c) a conserved core microbial communities are associated with olive tree's root endosphere and soil rhizosphere zones. The Illumina-based amplicon sequencing helped us characterize and compare the size and structure of olive tree-associated bacterial and fungal communities, providing comprehensive insights into their relevance in non-traditional production areas.

The trees of the cultivar Arbequina grown at the three geographically distinct locations (Carrizo Springs, Moulton, Berclair) were selected to analyze the microbiome profile of olive rhizosphere soil and endosphere of roots over three seasons (Spring, summer, and fall). Amplicon sequencing of 16S rRNA and ITS regions on the Illumina Miseq platform generated 8,971,015 bacterial and 8,049,726 fungal raw sequence reads. After filtering, 8,879,079 bacterial and 6,868,796 fungal high-quality sequence reads were obtained.

Effects of location and seasons on OTUs

The number of OTUs was used to provide a comprehensive overview of the microbial structure and distribution, where 14190 bacterial OTUs were identified in the rhizosphere (RS) and root endosphere (RE) samples. A Venn diagram representing OTU distribution in the rhizosphere and root endosphere of all samples revealed that 73.57% (10,441 OTUs) were only found in the rhizosphere soil as opposed to the roots (26.12%, 3707 OTUs), and both niche samples shared 0.03% (5 OTUs) of the total bacterial OTUs. Of all locations, rhizosphere soils shared 43.81% (4575 OTUs), while roots endosphere shared 43.78% (1623 OTUs of total bacterial OTUs). The most unique bacterial OTUs were found in rhizosphere soils (14.13%, 1476 OTUs) in the Carrizo Springs and Moulton endosphere roots (17.56%, 651 OTUs). Across seasons, 5277 OTUs (50.54%) and 1895 OTUs (51.11%) of rhizosphere and root endosphere OTUs were shared between seasons, where1176 OTUs in spring (11.26%) and 639 OTUs in fall (17.23%) exhibited the unique rhizosphere and root endosphere OTUs (Fig. 1; Fig.S1). In the case of fungi, 11068 OTUs were generated. While both niches shared 7.17% (794 OTUs) of the total fungal OTUs, 94.71% (10483 OTUs) were exclusively detected in rhizosphere soil compared to roots (5.29%, 585 OTUs). In Berclair, rhizosphere soils (30.5%, 2817 OTUs) and endosphere (22.57%,195 OTUs) exhibited the most unique fungal OTUs. The rhizosphere soils of all locations shared 21.29% (1966 OTUs), whereas the endosphere roots shared 33.62% (2786 OTUs). However, for seasons, spring (27.81%,2304 OTUs) and summer (29.37%, 220 OTUs) displayed the unique OTUs for rhizosphere soils and root endosphere, respectively. Similarly, the rhizosphere and root endosphere shared 25.16% (2,786 OTUs) and 41.35% (528 OTUs) OTUs between seasons (Fig.S2).

Location and seasons structure the microbial community composition

The relative abundance of dominant bacterial phyla varied across all locations and seasons for both sample categories. In the analysis, RS is an acronym for rhizosphere soil, RE for root endosphere, CS for Carrizo Springs, M for Moulton, B for Berclair, Sp for spring, Su for summer, and F for fall. Phylum Proteobacteria (RS, 16.42%; RE, 15.498%) and Actinobacteria (RS, 12.63%; RE, 16.47%) dominated the bacterial communities in both the rhizosphere and endosphere (Fig. 1, Table S1) followed by Proteobacteria (RS, 16.42%; RE, 15.498%) and Firmicutes (RS, 4.877%; RE, 11.1124.25%). The archaeal phylum Crenarcheota was found only in the rhizosphere (4.92%), while it showed zero abundance in the endosphere. At the genus level, Bacillus was identified as the dominant taxon across niches (RS, 3.41%; RE, 5.78%), followed by Rubrobacter (5.27%) in the rhizosphere. Actinophytocola (3.49%), Unidentified Streptosporangiales (2.38%), and Pseudonocardia (1.6%) were identified only in the root endosphere. Intriguingly, a greater proportion of "Others" under phyla and genus was discovered in both niches, indicating the possibility of a diverse bacterial composition (Fig. 1A, Fig.S3A).

Considering the influence of locations within seasons for given rhizosphere soil niches for taxonomic composition, The top known predominant bacterial phyla were Actinobacteriota in Moulton in all seasons (Sp,15.38%; Su,14.19%; F,14.06%), Proteobacteria in Carrizo Springs summer and fall (Su,10.52%; F,7.62%) and Moulton summer (7.01%), and Firmicutes (Sp,6.45%; F,6.84%), archaeal phyla Crenarcheota (Sp,8.65%; Su,5.66%) were higher in Berclair location. In the root endosphere, there was an abundance of Actinobacteriota in Carrizo Springs in all seasons (Sp,17.66%; Su,19.88%; F,33.54%) while Proteobacteria was greater in Carrizo Springs summer (23.88%) and Berclair in all seasons respectively (Sp,23.02%; Su,16.96%; F,16.38%). The summer and fall of Berclair were also enriched with other phyla like Bacteroidota (Su,4.52%; F,6.81%), while Firmicutes was found in higher numbers in the spring and summer seasons of Moulton (Sp,17.87%; Su,18.82%) (Fig.S3B-C). At the genus level, Rubrobacter was dominant in the rhizosphere of Berclair in the spring and fall (8.8%) seasons, followed by Bacillus in the spring and fall of Berclair (4.64%) and Carrizo Springs (4.86%). The Berclair and Carrizo Springs rhizosphere was also enriched with Candidatus_Nitrosophaera (3.40%) in spring and fall. In contrast, the root endosphere of Carrizo Springs was enriched with Actinophytocola (F,11.19%), Cronobacter (Su,9.65%), and Unidentified Streptosporangiales (Sp,3.04%; Su,7.08%; F,10.01%). Berclair endosphere was abundant in Corynebacterium in spring (8.39%) (Fig. 1B-C).

Like bacteria, the relative abundance of fungal phyla varied across all geographic regions and climatic seasons in both niches. Phylum Ascomycota (RS,71.09%; RE,41.37%) dominated the fungi communities, which was the most abundant phylum in both the rhizosphere and endosphere (Fig. 1, Table S1), followed by Basidiomycota (RS, 10.96%; RE, 30.95%). While Mortierellomycota was exclusively identified in rhizosphere soil (3.60%), Glomeromycota was found in abundance in the root endosphere (7.72%). At the genus level, in addition to "Others" found across both niches, members of Chaetomiaceae were identified as the dominant taxon across the rhizosphere (RS,17.61%; RE,1.52%), while unidentified Agaricales (17.41%) and Xylariales (15.97%) were dominant in the root endosphere. Furthermore, a higher percentage of "Others" under the order, family, and genus was found in both niches, suggesting a potential for a diverse fungal composition (Fig. 2A, Fig. S4A). The top four abundant fungal phyla in all rhizosphere niches of locations within seasons were Ascomycota, with the highest in Berclair during fall (79.25%), in spring (79.71%) at Carrizo Springs and in fall (78.5% at Moulton. Basidiomycota came next during summer in Moulton (23.11%) and Berclair (14.89%) and during spring at Carrizo Springs (15.4%). Mortierellomycota was more prevalent in Berclair during spring (7.47%) and summer (7.19%). The phylum Glomeromycota was equally abundant in Carrizo Spring's summer and fall seasons (Su,6.10; F,3.52%). Phylum Ascomycota dominated the root endosphere and was particularly abundant in all locations in springtime (CZ,70.90%; M,63.2%; B,63.54%), followed by Basidiomycota in the Berclair (58.91%) and at Carrizo Springs in summer (35.58%) and fall (38.96%) seasons. Glomeromycota showed the highest abundance in summer of all locations (CZ,25.81%; M,11.12%; B,10.86%) (Fig. S4B-C).

At the family level, members of Chaetomiaceae were enriched in all seasons of Berclair (Sp,32.87; Su,11.83%, F,24.63%), Hypocrealeas were dominant in the rhizosphere of Moulton summer (24.32%), Pleosporaceae (Sp,18.02%) and Cucurbitariaceae (Su,21.34%) in Carrizo Springs, while Berclair rhizosphere was enriched with Mortierellaceae during spring and summer (Sp,7.47%; Su,7.19%). In contrast, in the root endosphere, Unidentified Xylariales dominated the spring of Berclair (49.85%) and Moulton (48.21%), Moulton root endosphere was found to be abundant in Herpotrichilleaceae during spring and summer (Sp,16.29%; Su,13.53%) and Glomeraceae was enriched in Carrizo Springs root in summer (25.77%) (Fig. 2B-C). Using linear discriminant analysis of effect size (LEfSe), we aimed to identify high dimensional biomarkers that most likely explained observed differences between locations or seasons. The linear discriminant analysis effect size (LEfSe) with logarithmic LDA > 2 was used to identify biomarker taxa. Overall, rhizosphere soil contained a more considerable number of bacterial biomarkers than endosphere soil (Fig.S5A). LEfSe analysis indicated that bacterial genera Rubrobacter, Candidatus_Nitrosophaera, and unidentified Nitrosophaeraceae were enriched in the rhizosphere. Root endosphere showed a significant presence of many genera, such as Bacillus, Actinophytocola, Streptosprangiales, and Pseudonocardia. Between locations, Candidatus _Nitrosophaera and members of Nitrosophaeraceae were the predominant rhizosphere soil genus in Berclair. Carrizo Springs was enriched with Sphingomonas, unidentified Geminicoccaceae, and Actinomarinales, while the Moulton rhizosphere contained Microtrichiales and Actinobacteria. In contrast, the root endosphere of Berclair was enriched with abundant genera such as Corynebacterium, Niastella, unidentified Micromonosporaceae, Rhizobium, and Promicromonospora. In comparison, Carrizo Springs and Moulton were enriched with only a few genera, such as Streptosporangiales Clostridium_senso_stricto_6 in Carrizo Springs and Exiguobacteium in Moulton (Fig. 3A, C).

Concerning fungal biomarkers, the rhizosphere was enriched with genera such as Mortierella, Aspergillus, Pyrenochaeta, Trichoderma, Acrophialophora, Humicola, Dothiora and unidentified Chaetomiaceae, Hypocreales and Agaricales, root endosphere was enriched with genera such as Penicillium, Subulicystidium, Pocescoma and unidentified members of Agaricomycetes, Xylanases, Auriculariales, and Cerataobasidicae (Fig.S6A). In rhizosphere soils of locations, besides members of Chaetomiaceae, Penicillium, Mortierella, Leucocoprinus, Humicola, Wilcoxina, and Rhexothecium were abundant in Berclair. Moulton carried members of Hypocreales, Pleosporales, Aspergillus, Trichoderma, Acrophailophora, and Curvularia. While no unique distinguishable biomarker fungi existed in Carrizo Springs, it contained moderate numbers of Hypocreales, Aspergillus, Curvularia, and Herpotrichellaceae. In contrast, the genus Penicillium was a prominent biomarker, followed by Mortierella and unidentified members of Ceratobasidiaceae, Micrascaceae, and Sordariomycetes in the root endosphere of Carrizo springs. On the other hand, the Moulton and Berclair root endospheres were frequently abundant with species from the genera Malassezia and Serendipitaceae, respectively (Fig. 3B, D).

No significant features were identified that distinguished the biomarkers among the three seasons in both the bacteria and fungi. We also examined the existence of distinct biomarkers between niches for each season. Spring showed enriched bacterial biomarkers in the rhizosphere soil, while summer showed more dominant biomarkers in the root endosphere than in other seasons. Genera like Rubrobacter, Nitrosophaeraceae, and Gaiellales genera were detected in all seasons in the rhizosphere during the spring and fall. While Microvirga was identified as a significant biomarker in soils in the summer and fall, Candidatus _Nitrosophaera was predominant in the spring and fall, and class α-Proteobacteria members were found in summer rhizosphere soil. The root endosphere of all seasons contained the genus Actinophytocola. Staphylococcus, Pseudonocardia, and Kibdelosporangium were exclusively identified in spring, Bacillus, Cronobacter, and Streptomyces in summer, and Saccharimonadales in fall. The root endosphere also included Rhizobium and Niastella during spring and summer and Streptosporangiales in the summer and fall (Fig.S5 B-D). Seasonally, fall demonstrated a greater concentration of enriched fungal biomarkers in the rhizosphere soil than in other seasons. While Mortierella, Aspergillus, Penicillium, Trichoderma, and Acrophialophora were found in the rhizosphere throughout all seasons. Wilcoxina, Clonostachys, and Acremonium were exclusively seen in the fall, and Pyrenocheta was only detected in the summer. While the genus Pyrenechaeota was only detected in the summer, the fall showed biomarkers like Trichoderma and Humicola. The predominant biomarkers in the root endosphere genera include Malassezia and members of Auriculariales in the spring, respectively, whereas Xylariales were found across all seasons. (Fig.S6B-D).

Bacterial and fungal taxonomic richness and diversity

Alpha diversity

Rarefaction curves demonstrated distinct differences in bacterial and fungal community composition as sampling depth increased for each sample from three locations in three seasons, indicating adequate sequencing effort and the total diversity within the sample was captured. With the increase in sample size, the Specaccum (species cumulative curve), like the rarefaction curves, showed the rate of increase of new species of bacteria and fungi (Fig. S7).

To verify its validity, alpha diversity metrics like Shannon (H') and Inverse Simpson (1/D) indices were used to assess the diversity of bacterial and fungal communities within samples across niches, locations, and seasons. Bacterial communities differed significantly (Kruskal-Wallis, P < 0.001) in diversity and species richness among overall sample categories, with rhizosphere soil exhibiting the highest diversity based on all metrics (H' = 6.27; 1/D = 162.61) (Fig. 4A, Fig.S8A). Season and site significantly affect the diversity of bacterial communities. For example, the site explained 83.91%, and the season explained 78.45% of the significant variation in bacterial diversity in the rhizosphere and root endosphere, respectively. The indices estimated a significant difference (Kruskal-Wallis, P < 0.05) in niches between locations for bacterial diversity and richness in rhizosphere soil. Moulton rhizosphere soil exhibited much higher richness and diversity than other locations (H’=6.45). In contrast, none of the used metrics revealed a significant difference in the α-diversity of the root endosphere niche in locations (Table S1; Fig. 4C). For rhizosphere soils, seasonal differences in bacterial diversity and richness were not statistically significant, whereas in root endosphere, seasonal differences in α -a significant variation in observed richness can explain diversity. At the same time, the evenness index comparison (InvSimpson) was not statistically significant (Kruskal-Wallis, P > 0.05). In particular, the root endosphere had a greater bacterial diversity in the fall season (H' = 6.27) than in the other seasons based on the diversity metrics used in the study (Fig. 4E, 4h; Table S1; Fig.S8A, C, E). Fungi communities differed considerably (Kruskal-Wallis, P < 0.001) in diversity and species richness between niches, with rhizosphere soil showing the highest diversity across all locations and seasons (H’= 3.44; 1/D = 13.06) (Table S2; Fig. 4B; S8B). However, no statistically significant variation in fungal richness and evenness in the rhizosphere and root endosphere was observed according to the metrics used (Kruskal-Wallis, P > 0.05) (Fig. 4D, F; S8D, F).

Beta diversity: Permutational multivariate analyses of variance (PERMANOVA) of the Bray-Curtis distance matrix showed that compartments (R²; Bacteria:0.451; Fungi:0.123) significantly influenced microbial community differences (P = 0.001). Furthermore, locations in the rhizosphere (R²; Bacteria:0.305, Fungi:0.216) and endosphere (R²; Bacteria:0.146, Fungi:0.156) influenced microbial diversity. Seasons did not affect bacterial diversity in either niche; only fungal diversity was found in the rhizosphere (R² = 0.084, P = 0.001). The principal coordinate analysis (PCoA) based on Bray-Curtis's distance revealed that the 54 samples of the rhizosphere and endosphere bacterial communities from the same plant species exhibited a clear tendency to group based on sample type, with two principal component scores accounting for 45.7% and 4.3% of the total variations for bacteria and 14.8% and 7.0% of the total variations for fungi, respectively (Fig. 5). Rhizosphere bacterial samples from Berclair and Moulton tend to have separate clusters and Carrizo Springs formed separate clusters from other sites. There was an overlapping clustering for the root endosphere of Berclair and Moulton, while samples from Carrizo Springs within seasons were scattered (Fig. 5C, E). Carrizo Springs' rhizosphere soil samples for fungi were dispersed for all seasons, while Moulton and Berclair samples overlapped. For root endosphere samples, samples did not cluster clearly between Berclair and Moulton locations, while Carrizo Springs clustered from other locations, although not season-wise, because they overlapped (Fig. 5D, F). These differences were also shown by NMDS in which clear separation of samples according to the locations was observed.

In contrast, samples based on seasons displayed higher overlapping for bacteria and fungi (Fig.S9). Comparable results were observed in the Bray-Curtis dissimilarity distance-based hierarchical cluster analysis, demonstrating that the samples clustered into distinct groups based on the species composition of each sample. Rhizosphere samples were grouped separately from root endosphere samples. Based on locations and seasons, the two groups were even further subdivided. Based on locations, for bacteria, it was evident that the rhizosphere soil samples from Carrizo Springs and Moulton clustered with those from Berclair, whereas the root endosphere samples from Berclair and Moulton clustered within Carrizo Springs. Likewise, root endosphere samples of fall and spring were located within summer, which agreed with PCoA and NMDS analyses. As for fungi, the rhizosphere soil and root endosphere samples from Moulton and Berclair strongly clustered separately with those from Carrizo Springs based on their geographic locations. The rhizosphere samples from the fall and spring were grouped within the summer, whereas the root endosphere samples from the seasons were intermingled (Fig.S10).

Plant—associated microbial communities are critical determinants of plant health and productivity, contributing to nutrient availability and enhancing tolerance to abiotic and biotic stress [38–40]. With changing dynamics of the climate-driven amplification of abiotic stresses such as drought and heat, global olive cultivation has become highly vulnerable to sudden outbreaks of new diseases or herbivorous insect pests [41]. Unlike seasonal crops, fruit or nut trees like olives have a fundamentally different relationship with the soil. The olive-associated microbiome has been identified as a rich source of microbes that exhibit promise as agents that promote plant growth and exert biocontrol effects [42]. This study aimed to characterize the bacterial community compositions of the rhizosphere and endosphere of the Arbequina olive cultivar grown in three geographical regions from spring to fall. Numerous environmental and host-related factors, such as geographic location, plant genotype and phenotype, soil chemistry, and seasonal influences, will likely impact the microbial communities linked to plant hosts [43]. Our results showed a higher number of rhizospheres OTUs and, instead, a small number of roots endosphere OTUs for bacteria and mycobiota (Fig. S4). This result aligns with other studies where the microbial communities from the olive root endosphere are less diverse than those from the rhizosphere [20]. The number of shared and unique OTUs suggests that rhizosphere samples contained most OTUs in both bacterial and fungal datasets, confirming that soil serves as a primary reservoir for potential root endophytes [44]

The rhizosphere strongly influences the richness and diversity of microbial communities:

Overall, Alpha diversity analysis revealed that compared to roots, the rhizosphere soil had a significantly higher level of microbial diversity, presumably because the root endosphere tends to create an inner environment that is relatively stable, resulting in fewer changes in the microbial community within the plant. Locations specifically affected the rhizosphere's bacterial abundance and fungal richness but did not impact the root endosphere's community composition. Moulton and Berclair demonstrated the highest diversity for bacterial evenness and fungal richness in community composition. Recent studies in several experimental systems have found that fungal communities are more spatially differentiated than prokaryotic communities [45–47], suggesting that fungal endemism may shape communities at multiple scales and habitats.

Contrary to other research findings, which indicated that the alpha diversity of the rhizo-biome increased between seasons [48], the seasons increased both richness and evenness for bacteria in the root endosphere. The greater alpha diversity in summer and community compositional differences between spring and summer suggest that root-associated microbiota alterations are linked to plant phenological processes. In addition to indirectly affecting microbes, climatic conditions also influence the rate of photosynthesis and, consequently, the rate of rhizodeposition, as demonstrated for trees and perennial plants [49]. Thus, seasonal changes in bacterial communities are most likely influenced by an enhanced carbon flux from increasing temperatures [50]. Together, these findings imply that the temporal dynamics in the root-associated microbiota are affected by plant phenology and abiotic factors like weather, which may have an immediate impact on the microbiota or may have an indirect effect by altering plant physiological processes. However, seasons did not affect fungal alpha diversity in either compartment. Further research is needed to determine if these disparities are due to nutrient availability in different soils and sample types (rhizosphere vs. root endosphere) or the trophic characteristics of the bacterial and fungal species.

Geographic location influences the beta diversity of microbial communities.

In the present study, the beta-diversity analysis revealed that rhizospheric compartments and geographical location were the primary drivers of the compositional variations of microbial communities. Comparing the rhizosphere to the endosphere, we observed a greater diversity of fungi in the rhizosphere. Variations in bacterial composition were nearly the same. Because of this, as previously described in [51], our observations further imply that endophytic root colonization is not a passive process and that olive plants can choose from soil microbial consortia. Mature olive trees, like the one studied in this study, may have established endophytic microbiomes with mutualistic links to their hosts, resulting in less diversity [52]. Furthermore, a distinct clustering was seen amongst sample niches due to their geographical locations, indicating that the microbial populations that occupy olive tree niches differ spatially. Intriguingly, the rhizosphere samples from Carrizo Springs are the most dispersed in the ordination plot, indicating that the rhizospheres of these plants exhibit extremely varied composition and structure, even if the root endosphere of this location formed a distinct cluster. The overlapping clusters of the microbial communities from Moulton and Berclair, which are geographically closer (135 km apart), were generally more similar than those from distantly located Carrizo Springs (which is 341 and 285 km from Moulton and Berclair, respectively). As seasons did not affect microbiome variation in our study, variations in soil attributes, such as physicochemical composition, soil conductivity, and pH, could likely have a prominent role [53, 54].

Distinct microbial diversity associated with rhizocompartments across locations and seasons Microbial populations respond differently at all taxonomic levels, forming distinct soil microbial communities in soils with varying physicochemical characteristics [55]. Soil pH affects the bacterial component of soil microbial communities at continental, landscape, field, and levels, as bacteria have narrower pH ranges for optimal growth compared to fungi [56–59]. In our study, Actinobacteria and Proteobacteria dominated the rhizospheric and root endosphere bacterial community, consistent with previous studies [20, 52, 60] across all locations, with the former higher in the Moulton rhizosphere and the latter abundant in the root endosphere of Carrizo Springs and Berclair. Actinobacteria, the most abundant phylum in soil, produce extracellular enzymes, secondary metabolites (e.g., antimicrobial agents), and fast-degrading low-biodegradable organic compounds like hydrocarbons, lignin, and humus [61, 62]. Proteobacteria live in nutrient-rich soils and mineralize many soil nutrients [63]. In our study, Proteobacteria were observed during summer, as in Agave species during the dry season [64], and Actinobacteria in spring.

Additionally, the most prevalent genera of Actinobacteriota in the rhizosphere and root endosphere were Rubrobacter, Sphingomonas, Actinophytocola, Niastella, Pseudonocardia, and Unidentified Gaielleles and Streptosporangiales. While Rubrobacter and Sphingomonas colonize drought environments and metabolize pesticides and pollutants, Actinophytocola spp. in olive roots as endophyte can impact olive fitness and health. Our data derived from rhizospheric soil and root microbiome are consistent with the study [20] that examined root endophytic core microbiome in olive varieties and found Actinophytocola, Pseudonocardia, Bradyrhizobium to be essential for plant fitness. An abundance of Actinophytocola could benefit olives; endophytic Actinomycetes or their cultivated broths have been shown to inhibit the growth of plant pathogens [64]. The prevalence of Actinobacteria in our study (Rubrobacter, Actinophytocola, Pseudonocardia, unidentified Micromonosopraceae) in the root endophytic community suggests the possibility of isolating culturable representatives of these genera for their potential use as plant growth promotion (PGP) and biological control against olive tree pathogens. The rhizosphere of Berclair and Carrizo Springs and root endosphere of olives grown in Moulton were abundant in Phylum Firmicutes. Members of the genus Bacillus, well-known antagonists, and biocontrol agents are also the major components of the olive tree endosphere microbiota [65]. Synthesis of antimicrobial lipopeptide biosurfactants enabled this genus to be approved as a plant disease biocontrol agent [66–68]. The presence of other abundant phyla across both niches in all locations, such as Acidobacteriota, Bacterioidota, Myxococota, Gemmatimonadota, and Patescibacteria promote plant growth, protect against abiotic and biotic stresses, decompose soil organic matter, degrade plant-derived polymers, and involved in nutrient cycling [69–74]. No sequence reads from the kingdom Archaea were found in our investigation's root endosphere, as observed in the other studies [20]. However, the rhizosphere soil was enriched with Creanarcheota (now known as Thermoproteota). Members of this species are keystone members of agricultural soil communities due to their ability to promote the nitrogen cycle, fix carbon dioxide, and possess genes linked to plant growth promotion (PGP), suggesting their importance in these microbiomes [75]. The greater abundance of Candidatus_Nitrososphaeria in the rhizospheric soil of Berclair in the spring and fall seasons was an intriguing finding. As key players in the soil N cycle, Nitrososphaeria are frequent soil ammonia-oxidizing archaea [76]. A thaumarchaeal candidate genus Nitrososphaera as a core microbiome member was also identified in the endosphere of olive varieties [65] and associated soil [77].

Regarding fungi composition, both rhizocompartments have Ascomycota and Basidiomycota as the major fungi, especially in Moulton and Berclair, which is consistent with earlier research on the endophytic and rhizospheric communities of olive trees [20, 52]. Additionally, variations in the relative abundance of unidentified fungi, like Mortierellomycota and Glomeromycota, were discovered between compartments. Glomeromycota was more prevalent in the root endosphere of Carrizo Springs. Plants and Glomeromycota have an ancient and significant symbiosis [78]. Glomeromycota is also a monophyletic group of olive tree-dominant arbuscular mycorrhizal fungus (AMF) [79]. The AMF genera Rhizophagus, Glomus, and Gigaspora are known to improve host plant health by activating defense mechanisms against soilborne pathogens like Phytophthora, Fusarium, and Verticillium. Olive rhizosphere soils contain unusual phyla like Olpidiomycota, which can adapt to diverse environmental conditions, nutrition availability, and soil pH [80]. Berclair soil was enriched with Members of phyla Mortierellomycota, known to solubilize soil phosphorus and enhance available phosphorus [81]. The other phyla, such as Mucoromycota and Zoopagomycota, were rich in Carrizo Springs rhizosphere. Members of the Mucoromycota are beneficial or pathogenic, depending on where they are in the phylogenetic tree, where the latter taxon comprises species that are mainly mycoparasites [82].

The rhizosphere soils of all locations were enriched with Chaetomiaceae, while the root endospheres were abundant in Glomeraceae, unidentified members of Agaricales and Xylariales. Intriguingly, Carrizo Springs rhizosphere and endosphere were enriched with unidentified members of Auriculariales. The presence of members of the Agaricaceae family, as well as unidentified Agaricales and Auricualriales that belong to both Basidiomycota, is in line with studies that have found numerous Agaricomycetes acting as saprophytes, mutualists, and plant endophytes [78]. Furthermore, litter decomposition occurs primarily in the higher soil layers due to the activity of litter decomposers, which are found mainly in the order Agaricales (Basidiomycota) among these mushroom-forming fungi [83]. The members of the families Xylariales, Chaetomiaceae, Aspergillaceae, and Pleosporaceae are classified under the phylum Ascomycota. Due to their widespread distribution and ability to produce bioactive secondary metabolites with a variety of bioactivities, Xylariales are the most studied filamentous fungus group [84], while Chaetomiaceae members may play a part in defensive mutualism [85, 86].

This study also found seasonal variations in the rhizosphere and root endosphere fungal endophytes. Additionally, the fungal composition changed from late spring to fall. The most abundant endophytes, for example, were Xylariales, accounting for 33% of all isolates. Its relative abundance dropped to 11% by summer and 4% by fall. Many fungal species are predicted to colonize olive trees during the warm, humid spring and early summer months. Specific endophytes can eventually establish themselves, while others may decrease or disappear from the community. According to Martins et al. [23], this could account for decreased endophytic fungal diversity from late spring to autumn. However, throughout niches, the relative abundance of Agaricaceae and unidentified Agaricales members had significantly increased (up to 10% and 26%, respectively) in the summer and fall. The fungi community undergoes annual seasonal changes due to a successional process, which could be the cause of the observed variations [87]. In addition, it has been suggested that the recruitment of endophytic fungi may also occur due to interspecific competition among the fungi and alterations in the chemistry of plant tissues during the phenological growth stages of the tree [88].

Enrichment of specific microbial taxa in the rhizocompartments

Through a LeFse analysis, we identified distinct microbial taxa between overall niches and niche locations in the current study. For bacterial taxa, overall, the rhizosphere had an enrichment of Rubrobacter. This genus produces biopolymers such as polyhydroxyalkanoates (PHA), regarded as extremolytes designed to survive in saline and dry environments [89]. Between rhizospheres of locations, ammonia-oxidizing archaea of the family Nitrososphaeraceae and the most represented genus Candidatus Nitrososphaera, dominate the Berclair rhizosphere soils [90, 91]. Genus Sphingomonas, which produces phytohormones in addition to bioremediation, was abundant in Carrizo Springs [92]. As a prominent indicator in Moulton soils, unidentified Microtrichales were associated with gluconeogenesis, glycolysis, chlorophyll and porphyrin metabolism, transcription factors, and photosynthetic proteins [93].

In the root endosphere, Niastella, Promicromonospora, and Reyranella were found to be significantly abundant in Berclair and Moulton. Species of Niastella are antibiotic-resistant, metal-tolerant, and chitosan-hydrolyzing and were reported in the lateral roots of [94]. Promicromonospora has demonstrated its biocontrol function in resistance to Phytophthora capsici infection in pepper and resistance to smut in sugarcane [95, 96], and denitrification and bacterial wilt control in Reyranella [97]. Exiguobacterium, a bacterial genus classified as a distinct taxon in Moulton, is renowned for its widespread distribution owing to its ability to utilize a wide variety of polysaccharides and its resistance to environmental stress [98], whereas members of unidentified Streptosporangiales enriched in Carrizo Springs is known to produce antimicrobial compounds [38]. Regarding fungal biomarkers, overall, Unidentified Chaetomiaceae predominated in rhizosphere soil, while unidentified Xylariales dominated in the root endosphere. Due to their widespread distribution and bioactive secondary metabolites, Xylariales are the filamentous fungus group that has been investigated the most [84]. Between the rhizosphere soils of locations, unidentified Chaetomiaceae, Penicillium, Mortierella, Wilcoxina, and Humicola were the most prevalent taxa in Berclair compared to other places. Members of Unidentified Chaetomiaceae of phylum Ascomycota are a saprotrophic fungus found in soil and organic composts that degrades cellulose and other organic materials by producing lytic enzymes and can be a biocontrol agent [99–101]. Mortierella spp. has been hypothesized to suppress soilborne pathogens through competition or plant nutrient uptake (Miao et al., 2016). Other genera, like Penicillium spp., have been demonstrated to degrade polyethylene and induce resistance to the cucumber mosaic virus in Arabidopsis and tobacco plants [102, 103]. Ectomycorrhizal fungus Wilcoxina and halotolerant Humicola spp., which demonstrated antifungal, antibacterial, and antiproliferative activities [104], were also identified. Hypocreales predominated in Carrizo Springs and Moulton and displayed Pleosporales, Aspergillus, and Acrophialophora as unique biomarkers. Many soil ecosystems contain Hypocreales, which decompose straw residue in arable soils, inhibit plant pathogens, and stimulate plant growth and abiotic stress tolerance by interacting directly with roots [105, 106]. Numerous fungi belonging to the Pleosporales order have demonstrated resistance to low water potential and the capacity to stimulate the growth of plant seedlings. However, the most prevalent fungus in soil, Aspergillus, is predominantly involved with the degradation of organic matter [107]. The Acrophialophora genus was discovered to have endophytic strains that ameliorate drought stress and have antagonistic effects against plant diseases [108, 109]. On the other hand, Malassezia was enriched in the root endosphere of Moulton, Serendipitaceae, and Curvularia in Berclair, Penicillium, Ceratobasidiaceae, Microascaceae, and Mortierella in Carrizo Springs. Meanwhile, Serendipitaceae spp. can form a mutualistic symbiosis with crops, such as maize, cotton, switchgrass, barley, and wheat, and can supply nutrients and water to the host crop. Curvularia, as a root endophyte of phylum Ascomycota, has been reported in olives in soils with little nutrients [23, 110]. Malassezia globosa has been linked to plant ectoparasitic defense [111], and fungal populations within the Microascaceae have been demonstrated to degrade labile carbon in soil [112].

The present investigation provides a detailed characterization of the microbiome composition in the rhizosphere and root endosphere of the olive cultivar Arbequina. Rhizosphere microbial communities, characterized by their substantial richness and diversity, are more strongly associated with specific locations than endosphere communities. Furthermore, the species that exhibit higher abundance in the rhizosphere and endosphere may benefit plant growth and overall health. In conclusion, the presence of rhizocompartment and variations in geographic locations significantly impacted microbial populations across different geographical regions and seasons, with minimal influence observed from seasonal variations. Our findings highlight the need to consider the local microbial population, soil environment, seasons, and plant genotypes in future microbiome research. However, studying the microbiome in all plant compartments is necessary to provide the complete context of the complexity of interactions between the host plant and microorganisms. Further, by employing shotgun metagenomics and microbiome-driven isolation techniques to identify members of persistent common taxa, one can elucidate the functional potential of the microbiome associated with olive trees. This fundamental knowledge establishes the basis for further investigation, which will utilize possible microbial consortia to conduct synthetic, in vitro community-based evaluation of this assembly process and the functional roles of the olive-associated microbes.

Sample collection. Soil and root samples were collected from the olive groves located at three geographically distinct locations in Texas: Carrizo Springs/CZ (28° 31' 27.5556'' N, 99° 51' 30.7836'' W), Moulton/M (29° 34' 27.948'' N, 97° 8' 48.444'' W), and Berclair/B (28° 31' 31.404'' N, 97° 35' 7.332'' W). 'Arbequina,' the most grown variety in the United States, was selected for the study. The average age of trees was between 6–8 years old at each grove. The soil and root samples were collected in triplicates 4–7 inches deep within one meter from the trunk of the independent tress. The soil top layer (3 inches) was discarded for the rhizosphere soil sample collection. The soil rigidly attached to the roots was collected for rhizosphere analysis. Root samples were collected from the same samples to analyze root-associated microbial communities. The samples were collected during Spring/Sp (Mar-May), Summer/Su (June-Aug), and Fall/F (Sep-Nov) from each olive grove and transported to the Texas A&M AgriLife Research Center, Uvalde, Texas, in 2021 and stored at -80C until processing (Table 1).

Table 1

**Characteristics of the samples.** Abbreviated sample names collected in triplicates for each location and season for both niches are listed.
Sample Type	Season	Location	Abbreviated sample names
Rhizosphere soil	Spring	Carrizo Springs	RPC1-1, RPC2-2, RPC3-3
		Moulton	RPM1-19, RPM2-20, RPM3-21
		Berclair	RPB1-37, RPB2-38, RPB3-39
	Summer	Carrizo Springs	RUC1-4, RUC2-5, RUC3-6
		Moulton	RUM1-22, RUM2-23, RUM3-24
		Berclair	RUB1-40, RUB2-41, RUB3-42
	Fall	Carrizo Springs	RFC1-7, RFC2-8, RFC3-9
		Moulton	RFM1-25, RFM2-26, RFM3-27
		Berclair	RFB1-43, RFB2-44, RFB3-45
Root endosphere	Spring	Carrizo Springs	EPC1-10, RPC2-11, RPC3-12
		Moulton	EPM1-28, EPM2-29, EPM3-30
		Berclair	EPB1-46, EPB2-47, EPB3-48
	Summer	Carrizo Springs	EUC1-13, RPC2-14, RPC3-15
		Moulton	EUM1-31, EUM2-32, EUM3-33
		Berclair	EUB1-49, EUB2-50, EUB3-51
	Fall	Carrizo Springs	EFC1-16, RPC2-17, RPC3-18
		Moulton	EFM1-34, EFM2-35, EFM3-36
		Berclair	EFB1-52, EFB2-53, EFB3-54

Total DNA extraction. DNA from each soil and root sample was extracted using the PowerSoil® DNA Isolation Kits (MO BIO Laboratories, Carlsbad, CA, USA) and ZymoBIOMICS DNA Kit (Zymo Research, Irvine, CA, USA), following the manufacturer's recommendations. For the rhizosphere soil, samples were washed in phosphate-buffered saline (PBS) solution for 20 min, centrifuged at 2000×g for 5 min, and the remaining soil pellet was frozen in liquid nitrogen and stored at − 80°C. For the root endosphere DNA collection, after removal of adhering soil by shaking vigorously, roots were washed twice in PBS by shaking in 250 ml sterile flasks with 50 ml PBS for 20 min, sonicated (10 min of 25-s cycles at 3500 Hz), rinsed with sterile distilled water, flash-frozen in liquid nitrogen, and stored at − 80°C until extraction.

Sequencing, Filtering of Reads, and Assembly

The soil and root DNA samples were sequenced by the Novogene Corporation (USA) for microbial communities. In the case of rhizosphere soil for bacterial 16S analysis, the V3-V4 region was amplified using primers with barcodes 341F 5′- CCTACGGGAGGCAGCAG − 3′ and 806 R 5′- GGACTACHVGGGTWTCTAAT − 3′. and fungal ITS1 gene region was amplified using the ITS5-1737F (5’-GGAAGTAAAAGTCGTAACAAGG-3') /ITS2-2043R (5'- GCTGCGTTCTTCATCGATGC-3') primers. In the case of endosphere root samples, for bacterial 16S analysis, the V5-V7 region was amplified using barcoded primers 799F 5′- AACMGGATTAGATACCCKG − 3′ and 1193R 5′- ACGTCATCCCCACCTTCC − 3′. and fungal ITS1 gene region was amplified using the ITS1F-F (5'- CTTGGTCATTTAGAGGAAGTAA-3') /ITS1-1F-R (5'- GCTGCGTTCTTCATCGATGC-3') primers. The PCR products from each sample were pooled, end-repaired, A-tailed, and further ligated with Illumina adapters. Libraries were sequenced on a paired-end Illumina platform to generate 250bp paired-end raw reads. The library quality was checked with Qubit, real-time PCR for quantification, and bioanalyzer for size distribution detection. Quantified libraries will be pooled and sequenced on Illumina platforms according to the required effective library concentration and data amount. Paired-end reads were merged using FLASH (V1.2.7) [27], and the splicing sequences were called raw tags. Quality filtering on the raw tags was performed to obtain the high-quality clean tags [28] according to the QIIME (V1.7.0) quality control process. The tags were compared with the reference database (SILVA138) using the UCHIME algorithm (UCHIME Algorithm) [29]. Effective tags were obtained after the removal of chimeric [30].

Operational taxonomy unit (OTU) cluster, taxonomic annotation, and diversity analysis: Sequence analyses were performed by Uparse software (Uparse v7.0.10) [29] using all the effective tags. Sequences with ≥ 97% similarity were assigned to the same OTUs, and a representative sequence for each OTU was used for further annotation. For each representative sequence, QIIME (Version 1.7.0) in the Mothur method was performed against the SSUrRNA database of SILVA 138 database (version 132) [31] for species annotation at each taxonomic rank (threshold: 0.8∼1) [32] (kingdom, phylum, class, order, family, genus, and species). Statistical analysis and visualization of graphs were conducted in R studio v. 2023-06-16 [33, 34] unless stated otherwise. The microbial community analysis was carried out using the phyloseq R package, with the ASV tables and taxonomic classifications serving as the input dataset [35]. The dataset was rarefied by randomly selecting sequences with low read counts. Using ""ggrare"" from the ""ranacapa"" package, the rarefaction curves on species richness were computed [36]. The taxonomic composition was shown using plot bars. Using the rarefied dataset, the Kruskal Wallis test was performed to assess changes in alpha diversity according to the Shannon diversity (H'), Inverse Simpson (1/D) metrics, followed by post-hoc Dunn's testing for multiple pairwise comparisons at P < 0.05. To check for variations in community structure between sample groups, Bray-Curti's dissimilarity-based permutational analysis of variance (PERMANOVA, 999 permutations) along with the Adonis test was employed to evaluate the effect of factors (niches, locations, and seasons) on microbial composition. Principal coordinate analysis (PCoA), non-metric multidimensional scaling (NMDS), and dendrograms were used to visualize and compare microbial community structure between sample groups based on the Bray-Curtis distance matrix. The linear discriminant analysis (LDA) effect size (LEfSe) method implemented in MicrobiomeAnalyst [37] was employed to discern distinct biomarkers of bacteria and fungi underlying the observed microbiome differences between the locations and seasons. A threshold score of 2 and a significant α of 0.05 were applied to each feature to calculate its effect size.

Sequence Accession Numbers

The bacterial and fungal microbiome sequence data generated in this study are deposited in the National Center for Biotechnology Information (NCBI) under the BioProject numbers PRJNA1032045, PRJNA1031998, PRJNA1032109, and PRJNA1032141. Bacterial 16S datasets for endospheric root samples and rhizospheric soil samples are archived under the NCBI Sequence Read Archive (SRA) submissions SUB13925573 and SUB13908653, and the fungal ITS datasets for endospheric root samples and rhizospheric soil samples under the SRA submissions SUB13925585 and SUB13925588.

ACKNOWLEDGMENTS

We gratefully acknowledge the valuable support provided by the Texas Association of Olive Oil (TXAOO) and its member growers by providing access to their olive orchards and supporting the research activities. We sincerely appreciate the contributions of undergraduate and graduate students and research associates for their assistance in sample collection, which made this research possible.

AUTHOR CONTRIBUTIONS

V.J. conceived and designed the study. D.T., M.J., and A.M. conducted the experiments and collected the data. D.T. performed the statistical analysis. D.T. and V.J. drafted the manuscript. All authors approved the final version of the manuscript.

ADDITIONAL INFORMATION

The authors declare that they have no conﬂict of interest. All plant experiments were conducted in accordance with relevant institutional, national, and international guidelines and legislation.

FUNDING

We acknowledge the funding support through the Specialty Crop Block Grant program received for the Texas Association of Olive Oil by the Texas Department of Agriculture (Grant No. SC-1819-18) and the Hatch Program of the National Institute of Food and Agriculture, US Department of Agriculture [HATCH Project Accession No. 1011513; Project No. TEX09647].

DATA AVAILABILITY STATEMENT

The bacterial and fungal microbiome sequence data generated in this study are deposited in the National Center for Biotechnology Information (NCBI) under the BioProject numbers PRJNA1032045, PRJNA1031998, PRJNA1032109, and PRJNA1032141. Bacterial 16S datasets for endosphere root samples and rhizospheric soil samples are archived under the NCBI Sequence Read Archive (SRA) submissions SUB13925573 and SUB13908653, and the fungal ITS datasets for endosphere root samples and rhizospheric soil samples under the SRA submissions SUB13925585 and SUB13925588.

Zohary D, Spiegel-Roy P. Beginnings of Fruit Growing in the Old World: Olive, grape, date, and fig emerge as important Bronze Age additions to grain agriculture in the Near East. Science. 1975;187(4174):319–27.
Rallo L, Díez CM, Morales-Sillero A, Miho H, Priego-Capote F, Rallo P. Quality of olives: A focus on agricultural preharvest factors. Scientia horticulturae. 2018;233:491–509.
Rondanini D, Castro DN, Searles PS, Rousseaux MC. Fatty acid profiles of varietal virgin olive oils (Olea europaea L.) from mature orchards in warm arid valleys of Northwestern Argentina (La Rioja). Grasas y aceites. 2011;62(4):399–409.
Torres M, Pierantozzi P, Searles P, Rousseaux MC, García-Inza G, Miserere A, et al. Olive Cultivation in the Southern Hemisphere: Flowering, Water Requirements and Oil Quality Responses to New Crop Environments. Front Plant Sci. 2017;8; doi: 10.3389/fpls.2017.01830.
Ayerza R, Sibbett GS. Thermal adaptability of olive (Olea europaea L.) to the Arid Chaco of Argentina. Agriculture, ecosystems & environment. 2001;84(3):277–85.
Lanner RM. The Olive in California: History of an Immigrant Tree. Western Historical Quarterly. 2002;33(4):494–5; doi: 10.2307/4144779.
NASS. Noncitrus fruits and nuts 2021 Summary, National Agricultural Statistics Service (NASS) U.S. Department of Agriculture. https://www.nass.usda.gov/Publications/Todays_Reports/reports/ncit0522.pdf (2022). Accessed 2023 2022.
Malik* NSA, Bradford JM, Brockington J. Growing Olives in Texas. HortScience HortSci. 2004;39(4):799B-; doi: 10.21273/hortsci.39.4.799b.
TXAOO. 2021.
Tognetti R, d'Andria R, Sacchi R, Lavini A, Morelli G, Alvino A. Deficit irrigation affects seasonal changes in leaf physiology and oil quality of Olea europaea (cultivars Frantoio and Leccino). Annals of Applied Biology. 2007;150(2):169–86.
Di Vaio C, Nocerino S, Paduano A, Sacchi R. Influence of some environmental factors on drupe maturation and olive oil composition. Journal of the Science of Food and Agriculture. 2013;93(5):1134–9.
Mousavi S, de la Rosa R, Moukhli A, El Riachy M, Mariotti R, Torres M, et al. Plasticity of fruit and oil traits in olive among different environments. Scientific Reports. 2019;9(1):16968; doi: 10.1038/s41598-019-53169-3.
Parks S, Montague T. Influence of irrigation regime on gas exchange, growth, and oil quality of field grown, Texas (USA) olive trees. Open Agriculture. 2022;7:191–206; doi: 10.1515/opag-2022-0082.
Van Der Heijden MG, Bardgett RD, Van Straalen NM. The unseen majority: soil microbes as drivers of plant diversity and productivity in terrestrial ecosystems. Ecology letters. 2008;11(3):296–310.
Trivedi P, Leach JE, Tringe SG, Sa T, Singh BK. Plant–microbiome interactions: from community assembly to plant health. Nature Reviews Microbiology. 2020;18(11):607–21; doi: 10.1038/s41579-020-0412-1.
Scherber C, Eisenhauer N, Weisser WW, Schmid B, Voigt W, Fischer M, et al. Bottom-up effects of plant diversity on multitrophic interactions in a biodiversity experiment. Nature. 2010;468(7323):553–6.
Bizos G, Papatheodorou EM, Chatzistathis T, Ntalli N, Aschonitis VG, Monokrousos N. The Role of Microbial Inoculants on Plant Protection, Growth Stimulation, and Crop Productivity of the Olive Tree (Olea europea L.). Plants (Basel). 2020;9(6); doi: 10.3390/plants9060743.
Melloni R, Cardoso EJBN. Microbiome Associated with Olive Cultivation: A Review. Plants. 2023;12(4):897.
Fitzpatrick CR, Salas-González I, Conway JM, Finkel OM, Gilbert S, Russ D, et al. The Plant Microbiome: From Ecology to Reductionism and Beyond. Annu Rev Microbiol. 2020;74:81–100; doi: 10.1146/annurev-micro-022620-014327.
Fernández-González AJ, Villadas PJ, Gómez-Lama Cabanás C, Valverde-Corredor A, Belaj A, Mercado-Blanco J, et al. Defining the root endosphere and rhizosphere microbiomes from the World Olive Germplasm Collection. Scientific reports. 2019;9(1):20423.
Chialva M, De Rose S, Novero M, Lanfranco L, Bonfante P. Plant genotype and seasonality drive fine changes in olive root microbiota. Current Plant Biology. 2021;28:100219; doi: 10.1016/j.cpb.2021.100219.
Marasco R, Fusi M, Rolli E, Ettoumi B, Tambone F, Borin S, et al. Aridity modulates belowground bacterial community dynamics in olive tree. Environmental Microbiology. 2021;23(10):6275–91; doi: https://doi.org/10.1111/1462-2920.15764.
Martins F, Pereira JA, Bota P, Bento A, Baptista P. Fungal endophyte communities in above- and belowground olive tree organs and the effect of season and geographic location on their structures. Fungal Ecology. 2016;20:193–201; doi: https://doi.org/10.1016/j.funeco.2016.01.005.
de Oliveira AA, Ramalho MdO, Moreau CS, Campos AEdC, Harakava R, Bueno OC. Exploring the diversity and potential interactions of bacterial and fungal endophytes associated with different cultivars of olive (Olea europaea) in Brazil. Microbiological Research. 2022;263:127128; doi: https://doi.org/10.1016/j.micres.2022.127128.
Mina D, Pereira JA, Lino-Neto T, Baptista P. Impact of plant genotype and plant habitat in shaping bacterial pathobiome: a comparative study in olive tree. Scientific reports. 2020;10(1):3475.
Materatski P, Varanda C, Carvalho T, Dias AB, Campos MD, Rei F, et al. Spatial and temporal variation of fungal endophytic richness and diversity associated to the phyllosphere of olive cultivars. Fungal biology. 2019;123(1):66–76.
Magoč T, Salzberg SL. FLASH: fast length adjustment of short reads to improve genome assemblies. Bioinformatics. 2011;27(21):2957–63; doi: 10.1093/bioinformatics/btr507.
Bokulich NA, Subramanian S, Faith JJ, Gevers D, Gordon JI, Knight R, et al. Quality-filtering vastly improves diversity estimates from Illumina amplicon sequencing. Nature Methods. 2013;10(1):57–9; doi: 10.1038/nmeth.2276.
Edgar RC, Haas BJ, Clemente JC, Quince C, Knight R. UCHIME improves sensitivity and speed of chimera detection. Bioinformatics. 2011;27(16):2194–200; doi: 10.1093/bioinformatics/btr381.
Haas BJ, Gevers D, Earl AM, Feldgarden M, Ward DV, Giannoukos G, et al. Chimeric 16S rRNA sequence formation and detection in Sanger and 454-pyrosequenced PCR amplicons. Genome Res. 2011;21(3):494–504; doi: 10.1101/gr.112730.110.
Wang Q, Garrity GM, Tiedje JM, Cole JR. Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl Environ Microbiol. 2007;73(16):5261–7; doi: 10.1128/aem.00062-07.
Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 2013;41(Database issue):D590-6; doi: 10.1093/nar/gks1219.
Allaire J. RStudio: integrated development environment for R. Boston, MA. 2012;770(394):165–71.
Team RC. A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, 2021. Google Scholar There is no corresponding record for this reference.
McMurdie PJ, Holmes S. phyloseq: an R package for reproducible interactive analysis and graphics of microbiome census data. PLoS One. 2013;8(4):e61217.
Kandlikar GS, Gold ZJ, Cowen MC, Meyer RS, Freise AC, Kraft NJ, et al. ranacapa: An R package and Shiny web app to explore environmental DNA data with exploratory statistics and interactive visualizations. F1000Res. 2018;7.
Chong J, Liu P, Zhou G, Xia J. Using MicrobiomeAnalyst for comprehensive statistical, functional, and meta-analysis of microbiome data. Nature protocols. 2020;15(3):799–821.
Hardoim PR, Van Overbeek LS, Berg G, Pirttilä AM, Compant S, Campisano A, et al. The hidden world within plants: ecological and evolutionary considerations for defining functioning of microbial endophytes. Microbiology and molecular biology reviews. 2015;79(3):293–320.
Reinhold-Hurek B, Bünger W, Burbano CS, Sabale M, Hurek T. Roots shaping their microbiome: global hotspots for microbial activity. Annual review of phytopathology. 2015;53:403–24.
Jansson JK, Hofmockel KS. Soil microbiomes and climate change. Nature Reviews Microbiology. 2020;18(1):35–46.
Caselli A, Petacchi R. Climate change and major pests of Mediterranean olive orchards: are we ready to face the global heating? Insects. 2021;12(9):802.
Gharsallah H, Ksentini I, Frikha-Gargouri O, Hadj Taieb K, Ben Gharsa H, Schuster C, et al. Exploring Bacterial and Fungal Biodiversity in Eight Mediterranean Olive Orchards (Olea europaea L.) in Tunisia. Microorganisms. 2023;11(4):1086.
Coleman-Derr D, Desgarennes D, Fonseca‐Garcia C, Gross S, Clingenpeel S, Woyke T, et al. Plant compartment and biogeography affect microbiome composition in cultivated and native Agave species. New Phytologist. 2016;209(2):798–811.
Zarraonaindia I, Owens SM, Weisenhorn P, West K, Hampton-Marcell J, Lax S, et al. The Soil Microbiome Influences Grapevine-Associated Microbiota. mBio. 2015;6(2):10.1128/mbio.02527-14; doi: doi:10.1128/mbio.02527-14.
Peay KG, Kennedy PG, Talbot JM. Dimensions of biodiversity in the Earth mycobiome. Nature Reviews Microbiology. 2016;14(7):434–47; doi: 10.1038/nrmicro.2016.59.
Shakya M, Gottel N, Castro H, Yang ZK, Gunter L, Labbé J, et al. A Multifactor Analysis of Fungal and Bacterial Community Structure in the Root Microbiome of Mature Populus deltoides Trees. PLoS One. 2013;8(10):e76382; doi: 10.1371/journal.pone.0076382.
Meiser A, Bálint M, Schmitt I. Meta-analysis of deep-sequenced fungal communities indicates limited taxon sharing between studies and the presence of biogeographic patterns. New Phytologist. 2014;201(2):623–35; doi: https://doi.org/10.1111/nph.12532.
Ikeda S, Okazaki K, Takahashi H, Tsurumaru H, Minamisawa K. Seasonal Shifts in Bacterial Community Structures in the Lateral Root of Sugar Beet Grown in an Andosol Field in Japan. Microbes Environ. 2023;38(1); doi: 10.1264/jsme2.ME22071.
Amos B, Walters D. Maize root biomass and net rhizodeposited carbon: an analysis of the literature. Soil Science Society of America Journal. 2006;70(5):1489–503.
Becker MF, Hellmann M, Knief C. Spatio-temporal variation in the root-associated microbiota of orchard-grown apple trees. Environmental Microbiome. 2022;17(1):31; doi: 10.1186/s40793-022-00427-z.
Edwards J, Johnson C, Santos-Medellín C, Lurie E, Podishetty NK, Bhatnagar S, et al. Structure, variation, and assembly of the root-associated microbiomes of rice. Proc Natl Acad Sci U S A. 2015;112(8):E911-20; doi: 10.1073/pnas.1414592112.
Kakagianni M, Tsiknia M, Feka M, Vasileiadis S, Leontidou K, Kavroulakis N, et al. Above- and below-ground microbiome in the annual developmental cycle of two olive tree varieties. FEMS Microbes. 2023;4:xtad001; doi: 10.1093/femsmc/xtad001.
Szymańska S, Borruso L, Brusetti L, Hulisz P, Furtado B, Hrynkiewicz K. Bacterial microbiome of root-associated endophytes of Salicornia europaea in correspondence to different levels of salinity. Environ Sci Pollut Res Int. 2018;25(25):25420–31; doi: 10.1007/s11356-018-2530-0.
AlSharari SS, Galal FH, Seufi AM. Composition and Diversity of the Culturable Endophytic Community of Six Stress-Tolerant Dessert Plants Grown in Stressful Soil in a Hot Dry Desert Region. Journal of Fungi. 2022;8(3):241.
Franklin RB, Mills AL. Importance of spatially structured environmental heterogeneity in controlling microbial community composition at small spatial scales in an agricultural field. Soil Biology and Biochemistry. 2009;41(9):1833–40.
Fierer N, Jackson RB. The diversity and biogeography of soil bacterial communities. Proceedings of the National Academy of Sciences. 2006;103(3):626 – 31; doi: doi:10.1073/pnas.0507535103.
Lauber CL, Strickland MS, Bradford MA, Fierer N. The influence of soil properties on the structure of bacterial and fungal communities across land-use types. Soil Biology and Biochemistry. 2008;40(9):2407–15; doi: https://doi.org/10.1016/j.soilbio.2008.05.021.
Rousk J, Bååth E, Brookes PC, Lauber CL, Lozupone C, Caporaso JG, et al. Soil bacterial and fungal communities across a pH gradient in an arable soil. The ISME Journal. 2010;4(10):1340–51; doi: 10.1038/ismej.2010.58.
Deakin G, Tilston E, Bennett JM, Passey TAJ, Harrison NA, Fernández-Fernández F, et al. Spatial structuring of soil microbial communities in commercial apple orchards. Applied Soil Ecology. 2018;130:1–12.
Fausto C, Mininni AN, Sofo A, Crecchio C, Scagliola M, Dichio B, et al. Olive orchard microbiome: characterisation of bacterial communities in soil-plant compartments and their comparison between sustainable and conventional soil management systems. Plant Ecology & Diversity. 2018;11(5–6):597–610; doi: 10.1080/17550874.2019.1596172.
Olano C, García I, González A, Rodriguez M, Rozas D, Rubio J, et al. Activation and identification of five clusters for secondary metabolites in Streptomyces albus J1074. Microb Biotechnol. 2014;7(3):242–56; doi: 10.1111/1751-7915.12116.
Trinchera A, Migliore M, Warren Raffa D, Ommeslag S, Debode J, Shanmugam S, et al. Can multi-cropping affect soil microbial stoichiometry and functional diversity, decreasing potential soilborne pathogens? A study on European organic vegetable cropping systems. Frontiers in Plant Science. 2022;13; doi: 10.3389/fpls.2022.952910.
Fallah N, Yang Z, Tayyab M, Zhang C, Abubakar AY, Lin Z, et al. Depth-dependent influence of biochar application on the abundance and community structure of diazotrophic under sugarcane growth. PLoS One. 2021;16(7):e0253970; doi: 10.1371/journal.pone.0253970.
Matsumoto A, Takahashi Y. Endophytic actinomycetes: promising source of novel bioactive compounds. The Journal of Antibiotics. 2017;70(5):514–9; doi: 10.1038/ja.2017.20.
Müller H, Berg C, Landa BB, Auerbach A, Moissl-Eichinger C, Berg G. Plant genotype-specific archaeal and bacterial endophytes but similar Bacillus antagonists colonize Mediterranean olive trees. Frontiers in Microbiology. 2015;6; doi: 10.3389/fmicb.2015.00138.
Ben Abdallah D, Frikha-Gargouri O, Tounsi S. Bacillus amyloliquefaciens strain 32a as a source of lipopeptides for biocontrol of Agrobacterium tumefaciens strains. Journal of Applied Microbiology. 2015;119(1):196–207; doi: 10.1111/jam.12797.
Zicca S, De Bellis P, Masiello M, Saponari M, Saldarelli P, Boscia D, et al. Antagonistic activity of olive endophytic bacteria and of Bacillus spp. strains against Xylella fastidiosa. Microbiol Res. 2020;236:126467; doi: 10.1016/j.micres.2020.126467.
Cheffi M, Bouket AC, Alenezi FN, Luptakova L, Bełka M, Vallat A, et al. Olea europaea L. Root Endophyte Bacillus velezensis OEE1 Counteracts Oomycete and Fungal Harmful Pathogens and Harbours a Large Repertoire of Secreted and Volatile Metabolites and Beneficial Functional Genes. Microorganisms. 2019;7.
DeBruyn JM, Nixon LT, Fawaz MN, Johnson AM, Radosevich M. Global biogeography and quantitative seasonal dynamics of Gemmatimonadetes in soil. Appl Environ Microbiol. 2011;77(17):6295–300; doi: 10.1128/aem.05005-11.
Guo B, Zhang H, Liu Y, Chen J, Li J. Drought-resistant trait of different crop genotypes determines assembly patterns of soil and phyllosphere microbial communities. Microbiology Spectrum. 2023;11(5):e00068-23; doi: doi:10.1128/spectrum.00068-23.
Dedysh SN, Ivanova AA. Planctomycetes in boreal and subarctic wetlands: diversity patterns and potential ecological functions. FEMS Microbiology Ecology. 2018;95(2); doi: 10.1093/femsec/fiy227.
Ivanova AA, Zhelezova AD, Chernov TI, Dedysh SN. Linking ecology and systematics of acidobacteria: Distinct habitat preferences of the Acidobacteriia and Blastocatellia in tundra soils. PLoS One. 2020;15(3):e0230157; doi: 10.1371/journal.pone.0230157.
Larsbrink J, McKee LS. Bacteroidetes bacteria in the soil: Glycan acquisition, enzyme secretion, and gliding motility. Adv Appl Microbiol. 2020;110:63–98; doi: 10.1016/bs.aambs.2019.11.001.
Tian R, Ning D, He Z, Zhang P, Spencer SJ, Gao S, et al. Small and mighty: adaptation of superphylum Patescibacteria to groundwater environment drives their genome simplicity. Microbiome. 2020;8:1–15.
Nelkner J, Huang L, Lin TW, Schulz A, Osterholz B, Henke C, et al. Abundance, classification and genetic potential of Thaumarchaeota in metagenomes of European agricultural soils: a meta-analysis. Environmental Microbiome. 2023;18(1):26; doi: 10.1186/s40793-023-00479-9.
Kuypers MMM, Marchant HK, Kartal B. The microbial nitrogen-cycling network. Nature Reviews Microbiology. 2018;16(5):263–76; doi: 10.1038/nrmicro.2018.9.
Caliz J, Montes-Borrego M, Triadó-Margarit X, Metsis M, Landa BB, Casamayor EO. Influence of edaphic, climatic, and agronomic factors on the composition and abundance of nitrifying microorganisms in the rhizosphere of commercial olive crops. PLoS One. 2015;10(5):e0125787; doi: 10.1371/journal.pone.0125787.
Spatafora JW, Chang Y, Benny GL, Lazarus K, Smith ME, Berbee ML, et al. A phylum-level phylogenetic classification of zygomycete fungi based on genome-scale data. Mycologia. 2016;108(5):1028–46; doi: 10.3852/16-042.
Stürmer SL. A history of the taxonomy and systematics of arbuscular mycorrhizal fungi belonging to the phylum Glomeromycota. Mycorrhiza. 2012;22(4):247–58; doi: 10.1007/s00572-012-0432-4.
Idbella M, Bonanomi G. Uncovering the dark side of agriculture: How land use intensity shapes soil microbiome and increases potential plant pathogens. Applied Soil Ecology. 2023;192:105090; doi: https://doi.org/10.1016/j.apsoil.2023.105090.
Bao Z, Matsushita Y, Morimoto S, Hoshino YT, Suzuki C, Nagaoka K, et al. Decrease in fungal biodiversity along an available phosphorous gradient in arable Andosol soils in Japan. Can J Microbiol. 2013;59(6):368–73; doi: 10.1139/cjm-2012-0612.
Bonfante P, Venice F. Mucoromycota: going to the roots of plant-interacting fungi. Fungal Biology Reviews. 2020;34(2):100–13.
Floudas D, Bentzer J, Ahrén D, Johansson T, Persson P, Tunlid A. Uncovering the hidden diversity of litter-decomposition mechanisms in mushroom-forming fungi. The ISME journal. 2020;14(8):2046–59.
Becker K, Stadler M. Recent progress in biodiversity research on the Xylariales and their secondary metabolism. The Journal of Antibiotics. 2021;74(1):1–23.
Zimowska B, Nicoletti R. Arcopilus aureus: A valuable endophytic associate of hazelnut. Acta Agrobotanica. 2023;76:175998.
Nicoletti R, Di Vaio C, Cirillo C. Endophytic fungi of olive tree. Microorganisms. 2020;8(9):1321.
Wearn JA, Sutton BC, Morley NJ, Gange AC. Species and organ specificity of fungal endophytes in herbaceous grassland plants. Journal of Ecology. 2012;100(5):1085–92.
Fang W, Yang L, Zhu X, Zeng L, Li X. Seasonal and habitat dependent variations in culturable endophytes of Camellia sinensis. Journal of Plant Pathology & Microbiology. 2013;4(3):2157–471.
Kouřilová X, Schwarzerová J, Pernicová I, Sedlář K, Mrázová K, Krzyžánek V, et al. The first insight into polyhydroxyalkanoates accumulation in multi-extremophilic Rubrobacter xylanophilus and Rubrobacter spartanus. Microorganisms. 2021;9(5):909.
He J-Z, Hu H-W, Zhang L-M. Current insights into the autotrophic thaumarchaeal ammonia oxidation in acidic soils. Soil Biology and Biochemistry. 2012;55:146–54.
Zhalnina K, de Quadros PD, Gano KA, Davis-Richardson A, Fagen JR, Brown CT, et al. Ca. Nitrososphaera and Bradyrhizobium are inversely correlated and related to agricultural practices in long-term field experiments. Frontiers in microbiology. 2013;4:104.
Asaf S, Numan M, Khan AL, Al-Harrasi A. Sphingomonas: from diversity and genomics to functional role in environmental remediation and plant growth. Critical Reviews in Biotechnology. 2020;40(2):138–52.
Deng Y, Cheng C, Feng J, Liu S, Ma H, Chen X, et al. Rapid environmental change shapes pond water microbial community structure and function, affecting mud crab (Scylla paramamosain) survivability. Applied microbiology and biotechnology. 2020;104:2229–41.
Okazaki K, Tsurumaru H, Hashimoto M, Takahashi H, Okubo T, Ohwada T, et al. Community analysis-based screening of plant growth-promoting bacteria for sugar beet. Microbes and environments. 2021;36(2):ME20137.
Duan M, Wang L, Song X, Zhang X, Wang Z, Lei J, et al. Assessment of the rhizosphere fungi and bacteria recruited by sugarcane during smut invasion. Brazilian Journal of Microbiology. 2023;54(1):385–95.
Abbasi S, Spor A, Sadeghi A, Safaie N. Streptomyces strains modulate dynamics of soil bacterial communities and their efficacy in disease suppression caused by Phytophthora capsici. Scientific reports. 2021;11(1):9317.
Chen S, Qi G, Ma G, Zhao X. Biochar amendment controlled bacterial wilt through changing soil chemical properties and microbial community. Microbiological research. 2020;231:126373.
Zhang D, Zhu Z, Li Y, Li X, Guan Z, Zheng J. Comparative genomics of Exiguobacterium reveals what makes a cosmopolitan bacterium. Msystems. 2021;6(4):e00383-21.
Madbouly AK, Abdel-Wareth MT. The use of Chaetomium taxa as biocontrol agents. Recent Developments on Genus Chaetomium. 2020:251 – 66.
Silva AMM, Estrada-Bonilla GA, Lopes CM, Matteoli FP, Cotta SR, Feiler HP, et al. Does organomineral fertilizer combined with phosphate-solubilizing bacteria in sugarcane modulate soil microbial community and functions? Microbial ecology. 2022;84(2):539–55.
Soytong K, Kahonokmedhakul S, Song J, Tongon R. Chaetomium application in agriculture. Technology in Agriculture. 2021;229.
Elsharkawy MM, Shimizu M, Takahashi H, Ozaki K, Hyakumachi M. Induction of Systemic Resistance against Cucumber mosaic virus in Arabidopsis thaliana by Trichoderma asperellum SKT-1. Plant Pathol J. 2013;29(2):193–200; doi: 10.5423/ppj.Si.07.2012.01.
Ghosh SK, Pal S. De-polymerization of LDPE plastic by Penicillium simplicissimum isolated from municipality garbage plastic and identified by ITSs locus of rDNA. Vegetos. 2021;34(1):57–67.
Hosseyni Moghaddam MS, Safaie N, Soltani J, Pasdaran A. Endophytic association of bioactive and halotolerant Humicola fuscoatra with halophytic plants, and its capability of producing anthraquinone and anthranol derivatives. Antonie Van Leeuwenhoek. 2020;113(2):279–91; doi: 10.1007/s10482-019-01336-x.
Ma A, Zhuang X, Wu J, Cui M, Lv D, Liu C, et al. Ascomycota members dominate fungal communities during straw residue decomposition in arable soil. PLoS One. 2013;8(6):e66146.
Tyśkiewicz R, Nowak A, Ozimek E, Jaroszuk-Ściseł J. Trichoderma: The current status of its application in agriculture for the biocontrol of fungal phytopathogens and stimulation of plant growth. International Journal of Molecular Sciences. 2022;23(4):2329.
Nayak S, Samanta S, Mukherjee AK. Beneficial role of Aspergillus sp. in agricultural soil and environment. Frontiers in Soil and Environmental Microbiology. 2020:17–36.
Daroodi Z, Taheri P, Tarighi S. Direct antagonistic activity and tomato resistance induction of the endophytic fungus Acrophialophora jodhpurensis against Rhizoctonia solani. Biological Control. 2021;160:104696; doi: https://doi.org/10.1016/j.biocontrol.2021.104696.
Wang H, Wang Y, Kang C, Wang S, Zhang Y, Yang G, et al. Drought stress modifies the community structure of root-associated microbes that improve Atractylodes lancea growth and medicinal compound accumulation. Frontiers in Plant Science. 2022;13:1032480.
Ray P, Guo Y, Chi MH, Krom N, Saha MC, Craven KD. Serendipita bescii promotes winter wheat growth and modulates the host root transcriptome under phosphorus and nitrogen starvation. Environmental Microbiology. 2021;23(4):1876–88.
Renker C, Alphei J, Buscot F. Soil nematodes associated with the mammal pathogenic fungal genus Malassezia (Basidiomycota: Ustilaginomycetes) in Central European forests. Biology and Fertility of Soils. 2003;37(1):70–2; doi: 10.1007/s00374-002-0556-3.
Lueders T, Kindler R, Miltner A, Friedrich MW, Kaestner M. Identification of Bacterial Micropredators Distinctively Active in a Soil Microbial Food Web. Applied and Environmental Microbiology. 2006;72(8):5342–8; doi: doi:10.1128/AEM.00400-06.

No competing interests reported.

SupplementalFile1.pdf
Supplementary File 1: Tables S1 and S2.
SupplementalFile2.pdf
Supplementary File 2 Figures S1–S10.

Download PDF

Journal Publication

published 06 Aug, 2024

Read the published version in Scientific Reports →

Editorial decision: Revision requested
17 May, 2024
Reviews received at journal
12 May, 2024
Reviews received at journal
02 May, 2024
Reviewers agreed at journal
15 Apr, 2024
Reviewers agreed at journal
15 Apr, 2024
Reviewers invited by journal
13 Apr, 2024
Editor assigned by journal
05 Apr, 2024
Editor invited by journal
20 Mar, 2024
Submission checks completed at journal
20 Mar, 2024
First submitted to journal
10 Mar, 2024

You are reading this latest preprint version

Unraveling the Spatio-Temporal Dynamics of Soil and Root-Associated Microbiome in Texas Olive Orchards: A Comprehensive Analysis

Status:

Journal Publication

Version 1

Abstract

Figures

INTRODUCTION

RESULTS

Bacterial and fungal taxonomic richness and diversity

DISCUSSION

The rhizosphere strongly influences the richness and diversity of microbial communities:

Enrichment of specific microbial taxa in the rhizocompartments

MATERIALS AND METHODS

Declarations

References

Additional Declarations

Supplementary Files

Status:

Journal Publication

Version 1