ABSTRACT 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 of the olive-associated microbial

communities and specificity in two niches (rhizosphere and root endosphere), seasons (spring, summer, and fall), and domains (bacteria and fungi) in the microbiome of the olive cultivar

Arbequina across three olive orchards in Texas. Phylum Proteobacteria, followed by _Actinobacteriota_, dominated the bacterial populations in the rhizosphere and endosphere. _Rubrobacter_

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—in influencing

community differences. Microbial diversity was apparent within the endosphere and rhizosphere. The seasons influenced only the rhizosphere fungal diversity, contrasting the bacterial

diversity in either niche. The research provided a comprehensive overview of the microbial diversity in olive trees' rhizosphere and root endosphere. 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. SIMILAR CONTENT BEING VIEWED BY OTHERS COMPOSITIONAL

PROFILING OF THE RHIZOSPHERE MICROBIOME OF CANADA THISTLE REVEALS CONSISTENT PATTERNS ACROSS THE UNITED STATES NORTHERN GREAT PLAINS Article Open access 04 August 2024 SOIL BACTERIAL

COMMUNITY STRUCTURES IN RELATION TO DIFFERENT OIL PALM MANAGEMENT PRACTICES Article Open access 30 November 2020 SOIL FERTILITY IMPACT ON RECRUITMENT AND DIVERSITY OF THE SOIL MICROBIOME IN

SUB-HUMID TROPICAL PASTURES IN NORTHEASTERN BRAZIL Article Open access 16 February 2024 INTRODUCTION Olive (_Olea europaea_ L.) is cultivated commercially for the quality of its oil

worldwide. Although early domestication of olive trees began in the Mediterranean region1, in recent decades, commercial olive production expanded to non-traditional areas, such as Australia

and North and South America2, significantly varying in the agro-climatic conditions3,4,5. Olives were introduced into the United States in the late eighteenth century6. The United States

only represents less than 5% of the global olive production7. 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-1990s8, the high-density olive planting for oil production has been spread across approximately 1400 ha9. 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 climates10, the changing climate conditions have shown varied impacts

on olive fruit maturation and oil composition in different cultivars, locations, and water availability11,12,13. 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 productivity14. At the same time, plant-associated microbiomes confer fitness advantages to the plant host, including growth promotion, nutrient uptake, stress tolerance,

and resistance to pathogens15. In return, plants can affect soil microbial communities via host preference and changes in plant-derived inputs, such as litter, rhizodeposits, and root

exudates16. Several reviews have highlighted the significance of soil and olive tree-associated microbiomes in defining olive tree productivity17,18. However, since the Mediterranean basin

has been the world's production center for olives, research on the interactions with soil microbiota and/or olive tree microbiomes outside these non-traditional production areas is

lacking. Microbiota, particularly at the root level, are critical in modifying plant physiology and metabolism under various climatic conditions19. 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 soils20. Healthy and highly stable

root microbiota are critical in helping olive trees thrive in new environments and climatic conditions21. 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 rhizosphere20. 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 rainfall22. Likewise, the impact of

variables such as plant age, organ type, altitudinal gradient, geographic location, and season, but not the cultivar, on the structure of microbial communities in commercial olive plants

has been demonstrated23,24. On the contrary, studies have also shown the effect of the olive cultivar on the distinct differences in the endophytic and epiphytic microbial communities25,26.

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 (rhizosphere and root endosphere), seasons (Spring, summer, and

fall), and domains (bacteria and fungi) in the microbiome of the olive cultivar 'Arbequina,' the primary cultivar 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 rhizosphere communities

over the seasons or locations and (c) a conserved core microbial communities are associated with root endosphere and 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. RESULTS 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 14,190 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, 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, 1176 OTUs in spring (11.26%) and 639 OTUs in

fall (17.23%) exhibited the unique rhizosphere and root endosphere OTUs (Fig. 1; Supplementary Fig. S1). In the case of fungi, 11,068 OTUs were generated. While both niches shared 7.17% (794

OTUs) of the total fungal OTUs, 94.71% (10,483 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. For seasons, spring (27.81%, 2304 OTUs) and summer (29.37%, 220 OTUs) displayed the unique OTUs for rhizosphere soils

and root endosphere, respectively. (Supplementary Fig. S2). LOCATION AND SEASONS STRUCTURE THE MICROBIAL COMMUNITY COMPOSITION The relative abundance (R.A) of dominant bacterial phyla varied

across all locations and seasons for both sample categories. The analysis included the following acronyms: RS for rhizosphere, 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, 15.50%) and _Actinobacteria_ (RS, 12.63%; RE, 16.47%) dominated the bacterial

communities in both niches (12–16%). 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, while _Actinophytocola_ (3.49%) and Unidentified _Streptosporangiales_ (2.38%) were identified only in the root endosphere. In

addition, both niches were associated with a higher relative abundance of "Others" under phyla and genera, indicating the possibility of a diverse bacterial composition

(Supplementary Fig. S3A,D). Considering the influence of locations within seasons for given rhizosphere soil niches for taxonomic composition, the predominant bacterial phyla were

_Actinobacteriota_ in Moulton in all seasons, _Proteobacteria_ in Carrizo Springs, and archaeal phyla _Crenarcheota_ in Berclair location. In the root endosphere, there was an abundance of

_Actinobacteriota_ in Berclair during spring, Proteobacteria in Carrizo Springs, and Firmicutes relatively abundant in Moulton (Supplementary Fig. S3B,C, Supplementary Table S1). At the

genus level, _Rubrobacter_ was dominant in the rhizosphere of Berclair in the spring and fall seasons, followed by _Bacillus_ in the spring and fall of Carrizo Springs. In contrast, the root

endosphere of Carrizo Springs during Fall was enriched with _Actinophytocola_ (11.19%), while the Berclair endosphere was abundant in _Corynebacterium_ in spring (8.39%) (Fig. 1B,C,

Supplementary Table S1). In addition, both niches were associated with higher relative abundance of "Others" under phyla and genus, indicating the possibility of a diverse

bacterial composition. For Fungi, Phylum _Ascomycota_ dominated the fungi communities, the most abundant phylum in both niches (R.A > 40%). While _Mortierellomycota_ was exclusively found

in rhizosphere soil (3.60%), _Glomeromycota_ was found in higher abundance in the root endosphere (7.72%). At the family level, members of _Chaetomiaceae_ were identified as the dominant

taxon across the rhizosphere in all seasons, while unidentified _Agaricales_ and _Xylariales_ were abundant in the root endosphere (R.A > 10%). Furthermore, much like the bacterial

community, a higher percentage of "Others" under phyla and family was found in both niches, suggesting a potential for a diverse fungal composition (Fig. 2A, Supplementary Fig.

S4A,C). The top abundant fungal phyla in all rhizosphere niches of locations within seasons was _Ascomycota_ (R.A > 75%), followed by _Basidiomycota_. _Mortierellomycota_ was more

prevalent in Berclair during spring and summer (7%). Phylum _Ascomycota_ dominated the root endosphere and was particularly abundant in all locations in spring, while _Glomeromycota_ showed

the highest abundance in summer of all locations (Supplementary Fig. S4C,D, Supplementary Table S2). At the family level_,_ members of _Chaetomiaceae_ were observed in higher abundance in

all seasons of Berclair. Hypocrealeas in the rhizosphere of Moulton (24.32%) and _Cucurbitariaceae_ (21.34%) in Carrizo Springs were abundant during summer. In contrast, in the root

endosphere, in spring, members of unidentified _Xylariales_ dominated Berclair (49.85%) and Moulton (48.21%). In summer, the Moulton root endosphere was abundant in _Herpotrichilleaceae,_

and _Glomeraceae_ was abundant in Carrizo Springs roots (Su, 25.77%) (Fig. 2B, Supplementary Table S2). Using linear discriminant analysis of effect size (LEfSe), we aimed to identify

distinct taxa as biomarkers that most likely explained the differential relative abundance between locations or seasons. The linear discriminant analysis effect size (LEfSe) with logarithmic

LDA > 2 was used to identify markers. Overall, rhizosphere soil contained a more considerable number of bacterial markers than endosphere soil (Supplementary Fig. S5A–D). Between

locations, members of _Nitrosophaeraceae, Microtrichiales,_ and _Sphingomonas_ were the predominant rhizosphere soil genera in Berclair, Moulton, and Carrizo Springs, respectively. The root

endosphere revealed the presence of many distinct genera in Berclair (_Corynebacterium_, _Niastella, Rhizobium_, and _Promicromonospora)_ (Fig. 3A,C). Concerning fungal biomarkers, members

of _Hypocreales_ and _Cheatomiaceae_ were found in rhizosphere soils in Moulton and Berclair. While no unique distinguishable biomarker fungi existed in Carrizo Springs, they contained

moderate numbers of _Hypocreales_. In contrast, members of families such as _Ceratobasidiaceae_ and _Microascaceae_ were enriched in Carrizo Springs more than in other locations in the root

endosphere. (Fig. 3B,D). We also examined the existence of distinct biomarkers between niches for each season. All seasons showed a higher abundance of _Rubrobacter_ in rhizosphere soil_._

The root endosphere contained the genus _Actinophytocola_. Seasonally, spring and fall demonstrated greater enriched fungal biomarkers in the rhizosphere soil than in summer. Meanwhile,

_Mortierella_ was found in the rhizosphere throughout all seasons. _Wilcoxina_ was exclusively seen in the fall, and _Pyrenocheta_ was only detected in the summer. The predominant biomarkers

in the root endosphere genera include _Malassezia_ in the spring, respectively, whereas _Xylariales_ were found across all seasons. (Supplementary Fig. S6A–D). BACTERIAL AND FUNGAL

TAXONOMIC RICHNESS AND DIVERSITY ALPHA DIVERSITY Rarefaction curves indicated that the sequencing effort was sufficient to capture the total alpha diversity within the sample. With the

increase in sample size, the Specaccum (species cumulative curve) showed the rate of increase of new species of bacteria and fungi (Supplementary Fig. S7A–C). The alpha diversity metrics,

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, chi-squared, 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, Supplementary Fig. S8A). Season and site significantly affected the diversity of bacterial communities. For example, the

site and the season explained 83.91% and 78.45% of the significant variation in bacterial diversity in the rhizosphere and root endosphere, respectively. Among locations, Moulton rhizosphere

soil exhibited much higher richness and diversity than other locations (H′ = 6.45). None of the used metrics revealed a significant difference in the α-diversity of the root endosphere

niche in locations (Supplementary Table S3; Fig. 4C). For rhizosphere soils, seasonal differences in bacterial diversity and richness were not statistically significant. However, in the root

endosphere, seasonal differences showed a significant variation in observed richness, while the evenness index comparison (InvSimpson) was not statistically significant (Kruskal–Wallis,

chi-squared, P > 0.05). In particular, the root endosphere had a higher bacterial diversity in the fall season (H′ = 6.27) than in the other seasons (Fig. 4C; Supplementary Table S3;

Supplementary Fig. S8A,C,E). Fungi communities differed considerably (Kruskal–Wallis, chi-squared, P < 0.001) in diversity and species richness between overall niches, with rhizosphere

soil showing the highest diversity across all locations and seasons (H′ = 3.44; 1/D = 13.06) (Supplementary Table S4; Fig. 4B; Supplementary Fig. S8B). However, for location and season, no

statistically significant variation in fungal richness and evenness in the rhizosphere and root endosphere was observed according to the metrics used (Kruskal–Wallis, chi-squared, P > 

0.05) (Fig. 4D,F; Supplementary Fig. S8D,F). BETA DIVERSITY Permutational multivariate analyses of variance (PERMANOVA) of the Bray–Curtis distance matrix showed that compartments (_R_2;

Bacteria: 0.451; Fungi: 0.123) significantly influenced microbial community differences (P = 0.001). Furthermore, locations in the rhizosphere (_R_2; Bacteria:0.305, Fungi:0.216) and

endosphere (_R_2; 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

(_R_2 = 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), for fungi rhizosphere soil samples from Carrizo Springs 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 (Supplementary Fig. S9A–F). 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 (Supplementary Fig. S10A,B). DISCUSSION Plant-associated microbial communities are critical determinants of

plant health and productivity, contributing to nutrient availability and enhancing tolerance to abiotic and biotic stress27,28. 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 pests29. 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 effects30. 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 hosts31. Our results showed a higher number of rhizospheres OTUs and, instead, a small

number of roots endosphere OTUs for bacteria and mycobiota (Supplementary Fig. S4). This result aligns with other studies where the microbial communities from the olive root endosphere are

less diverse than those from the rhizosphere20. The number of shared and unique OTUs suggests that rhizosphere samples contained most OTUs in bacterial and fungal datasets, confirming that

rhizosphere soil is a primary reservoir for potential root endophytes32. THE RHIZOSPHERE STRONGLY INFLUENCES THE RICHNESS AND DIVERSITY OF MICROBIAL COMMUNITIE 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 bacterial abundance and fungal

richness but did not impact community composition in the root endosphere. Moulton and Berclair demonstrated the highest diversity in community composition for bacterial evenness and fungal

richness. Recent studies in several experimental systems have found that fungal communities are more spatially differentiated than prokaryotic communities33,34,35, 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 seasons36,

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 plants37. Thus, seasonal changes in bacterial communities are most likely influenced by

an enhanced carbon flux from increasing temperatures38. 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

in39, 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 diversity40. 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 role41,42. 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 characteristics43. In our study, _Actinobacteriota_ and _Proteobacteria_ dominated the rhizospheric and root endosphere bacterial community, consistent with previous

studies20,40,44 across all locations. _Actinobacteriota_, the most abundant phylum in soil, produces extracellular enzymes, secondary metabolites (e.g., antimicrobial agents), and

fast-degrading low-biodegradable organic compounds like hydrocarbons, lignin, and humus45,46. _Proteobacteria_ live in nutrient-rich soils and mineralize many soil nutrients47. Our study

observed _Proteobacteria_ during summer, as in _Agave_ species during the dry season48. Our data derived from rhizospheric soil and root microbiome are consistent with the study20 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, while _Rubrobacter_ colonizes drought environments and metabolizes pesticides and pollutants. The prevalence of _Actinobacteriota_ 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

microbiota49. Synthesis of antimicrobial lipopeptide biosurfactants enabled this genus to be approved as a plant disease biocontrol agent50. No sequence reads from the kingdom _Archaea_ were

found in our investigation in the root endosphere, as observed in the other studies20. However, the rhizosphere soil was enriched with _Creanarcheota_ (_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 microbiomes51. The greater abundance of soil ammonia-oxidizing archaea _Candidatus_Nitrososphaeria_ in the rhizospheric soil of Berclair in the

spring and fall seasons was an intriguing finding. A thaumarchaeal candidate genus _Nitrososphaera_ as a core microbiome member was also identified in the endosphere of olive varieties49 and

associated soil52. 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 trees20,40. Additionally, _Glomeromycota_ was more prevalent in the root endosphere of Carrizo Springs.

_Glomeromycota_ is a monophyletic group of olive tree-dominant arbuscular mycorrhizal fungus (AMF)53. 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_. Berclair soil was enriched with Members of phyla _Mortierellomycota_,

known to solubilize soil phosphorus and enhance available phosphorus54. The rhizosphere soils of all locations were enriched with _Chaetomiaceae_ members, while the root endospheres were

abundant in _Glomeraceae_, unidentified members of _Agaricales_ and _Xylariales_. While there were reports on bioactive metabolites produced by _Xylariales_55, _Chaetomiaceae_ members may

play a part in defensive mutualism56. 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 endophytes57. 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, which could account for decreased endophytic fungal diversity from late

spring to autumn23. 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 variations58. 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 tree59. ENRICHMENT OF SPECIFIC MICROBIAL TAXA IN THE RHIZOCOMPARTMENTS Through LeFse analysis, we identified distinct microbial taxa, such as Genus _Sphingomonas_, which

produces phytohormones and bioremediation, abundant in Carrizo Springs60. _Niastella_ have been described as metal-tolerant and chitosan-hydrolyzing61 in the root endosphere, whereas

members of unidentified _Streptosporangiales_ enriched in Carrizo Springs and during summer and fall seasons are known to produce antimicrobial compounds27. Regarding fungal biomarkers,

rhizosphere-enriched ectomycorrhizal _Wilcoxina_ and halotolerant _Humicola_ species have demonstrated antifungal, antibacterial, and antiproliferative activities62. _Hypocreales_ and

_Acrophialophora_ were unique biomarkers in Moulton and in Summer. Root endophytic _Hypocreales_ decompose straw residue in arable soils and support plant growth63, while _Acrophialophora_

and _Malassezia_ have been linked to alleviate drought stress and plant ectoparasitic defense64,65, _Serendipitaceae_ in Berclair, _spp_. can form a mutualistic symbiosis with crops and

supply nutrients and water to the host crop66. 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 communities 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 resident 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. MATERIALS AND METHODS 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 and 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 trees. 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 – 80 °C until processing (Table 1). 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. Unlike most studies, to minimize the plant-associated contamination of mitochondrial and chloroplast

sequences, we have deliberately opted to use different pairs of primers based on the pre-validated analysis in olives that resulted in higher capture of endospheric communities67,68. The

primers 799F/1193R showed the lowest mitochondria 16S rRNA amplification, no chloroplast sequences, and the highest numbers of bacterial OTUs68. Even if different primer sets for plant vs.

soil microbiomes have been shown to provide similar results for simultaneous studies on plant and soil microbiomes69, authors comprehend the likely bias introduced while comparing the

microbiomes across compartments due to the choice of different primers and suggest readers to restrict the taxonomic interpretations in the appropriate context and within specific

rhizocompartment. Despite the advantages, we agree that some differences in microbial communities could have resulted from the biased primer pairs selected. 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 250 bp paired-end raw reads. The

library quality was checked with Qubit, real-time PCR for quantification, and bioanalyzer for size distribution detection. Quantified libraries were 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)70, and the splicing sequences were called raw tags.

Quality filtering on the raw tags was performed to obtain the high-quality clean tags71 according to the QIIME (V1.7.0) quality control process. The tags were compared with the reference

database (SILVA138 and Unite V8.2 database) using the UCHIME algorithm72. Effective tags were obtained after the removal of chimeric sequences73. OPERATIONAL TAXONOMY UNIT (OTU) CLUSTER,

TAXONOMIC ANNOTATION, AND DIVERSITY ANALYSIS Sequence analyses were performed by Uparse software (Uparse v7.0.10)72 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) and the Mothur method were

performed against the SSUrRNA database of SILVA 138 with a threshold set to 0.8–174, blast with BLASTALL (Version 2.2.25) and Unite V8.2 database75 for species annotation at each taxonomic

rank (kingdom, phylum, class, order, family, genus, and species). Statistical analysis and visualization of graphs were conducted in R studio v. 2023-06-1676,77 unless stated otherwise. The

microbial community analysis was carried out using the phyloseq R package, with the OTU tables and taxonomic classifications serving as the input dataset78. 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 computed79. The taxonomic

composition was shown using plot bars. Using the rarefied dataset, the Kruskal Wallis chi-squared 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 dissimilarity matrix. The linear discriminant analysis (LDA) effect size (LEfSe) method implemented

in MicrobiomeAnalyst80 was employed to discern distinct biomarkers of bacteria and fungi underlying the observed microbiome differences between the locations and seasons. A threshold LDA

score of 2 and a significant α of 0.05 were applied to each feature to calculate its effect size. SEQUENCE ACCESSION NUMBERS The sequence data generated in this study are deposited in the

National Center for Biotechnology Information (NCBI) under the BioProject PRJNA1032045 (root endospheric) and PRJNA1031998 (soil rhizosphere) for bacterial and PRJNA1032109 (soil

rhizosphere), and PRJNA1032141(root endospheric) fungal microbiomes. DATA AVAILABILITY The bacterial and fungal microbiome sequence data generated in this study are deposited in the National

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  PubMed  Google Scholar  Download references ACKNOWLEDGEMENTS 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. 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]. AUTHOR INFORMATION AUTHORS AND AFFILIATIONS * Texas A&M AgriLife Research and Extension Center, Uvalde, TX, 78801, USA Dhivya

P. Thenappan, Dalton Thompson & Vijay Joshi * The University of Texas at San Antonio (UTSA), San Antonio, TX, 78249, USA Madhumita Joshi * Department of Botany, School of Life Sciences,

Mizoram University, Aizawl, 796004, India Amit Kumar Mishra * Department of Horticultural Sciences, Texas A&M University, College Station, TX, 77843, USA Vijay Joshi Authors * Dhivya P.

Thenappan View author publications You can also search for this author inPubMed Google Scholar * Dalton Thompson View author publications You can also search for this author inPubMed Google

Scholar * Madhumita Joshi View author publications You can also search for this author inPubMed Google Scholar * Amit Kumar Mishra View author publications You can also search for this

author inPubMed Google Scholar * Vijay Joshi View author publications You can also search for this author inPubMed Google Scholar 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. CORRESPONDING AUTHOR Correspondence to Vijay Joshi. ETHICS DECLARATIONS COMPETING INTERESTS The authors declare no competing interests. ADDITIONAL INFORMATION PUBLISHER'S

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Thenappan, D.P., Thompson, D., Joshi, M. _et al._ Unraveling the spatio-temporal dynamics of soil and root-associated microbiomes in Texas olive orchards. _Sci Rep_ 14, 18214 (2024).

https://doi.org/10.1038/s41598-024-68209-w Download citation * Received: 10 March 2024 * Accepted: 22 July 2024 * Published: 06 August 2024 * DOI: https://doi.org/10.1038/s41598-024-68209-w

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