“Ca. Nitrosocosmicus” members are the dominant archaea associated with plant rhizospheres
ABSTRACT
Archaea catalyzing the first step of nitrification in the rhizosphere possibly have an influence on plant growth and development. In this study, we found a distinct archaeal community, dominated by ammonia-oxidizing archaea (AOA), associated with the root system of pepper (Capsicum anuum L.) and ginseng plants (Panax ginseng C.A. Mey.) compared to bulk soil not penetrated by roots. While the abundance of total AOA decreased in the rhizosphere soils, AOA related to “Candidatus Nitrosocosmicus,” which harbor gene encoding manganese catalase (MnKat) in contrast to most other AOA, dominated the AOA community in the rhizosphere soils. For both plant species, the ratio of copy numbers of the AOA MnKat gene to the amoA gene (encoding the ammonia monooxygenase subunit A) was significantly higher in the rhizospheres than in bulk soils. In contrast to MnKat-negative strains from other AOA clades, the catalase activity of a representative isolate of “Ca. Nitrosocosmicus” was demonstrated. Members of this clade were enriched in H2O2-amended bulk soils, and constitutive expression of their MnKat gene was observed in both bulk and rhizosphere soils. Due to their abundance, “Ca. Nitrosocosmicus” members can be considered important players mediating the nitrification process in rhizospheres. The dominance of this MnKat-containing AOA in rhizospheres of agriculturally important plants hints at a previously overlooked AOA-plant interaction.
IMPORTANCE
Ammonia-oxidizing archaea (AOA) are widespread in terrestrial environments and outnumber other ammonia oxidizers in the rhizosphere, possibly exerting an influence on plant growth and development. However, little is known about the selection forces that shape their composition, functions, survival, and proliferation strategies in the rhizosphere. Here, we observed a distinct AOA community on root systems of two different plant species compared to bulk soil. Our results show that the “Ca. Nitrosocosmicus” clade, which possesses functional MnKat genes unlike most other AOA, dominated the rhizosphere soils. Moreover, members of this clade were enriched in H2O2-amended bulk soil, which mimics the ROS stress in root systems. While research on AOA-plant interactions in the rhizosphere is still in its infancy, these findings suggest that “Ca. Nitrosocosmicus” may be an important clade of AOA with potential AOA-plant interaction.
INTRODUCTION
Rhizosphere microbiome has been proposed to confer specific functions to its host plant, by modulating plant nutrient uptake, stress resistance, growth, and health (1–3). Soil types and characteristics are primarily shown to determine the background (bulk soil) microbiome, from which rhizosphere microbiomes are selected (4–9). Due to rhizodeposition, rhizospheres exhibit higher microbial abundances and distinct microbial communities compared to bulk soil (10–13).
Plant root exudates influence rhizosphere microbial community development by stimulating or inhibiting specific types of microorganisms (9, 14–16). Depending on the mode of photosynthesis (17) as well as the physiological and developmental status of the plant (18–20), roots release different types of exudates into the rhizosphere. Also, the phylogeny and genotype of a plant contribute to the development of distinct microbial communities in the plant rhizosphere by influencing the composition and activity of root exudates (13, 21–24). Conversely, it has been demonstrated that the rhizosphere microbiome affects root exudation inversely (2). Furthermore, it has been postulated that plants actively recruit soil microorganisms by releasing specific compounds into their rhizosphere that selectively stimulate specific microorganisms that are beneficial to plant growth and health (25–27). Signal molecules and antimicrobial compounds found in root exudates, such as phytoanticipins, phytoalexins, and sorgoleone, can be critical factors for shaping rhizosphere microbial communities (28–31). While we have gained a better understanding of the biology of root development as well as the structure and function of microbial communities in the rhizosphere, the interactions between rhizosphere microbiomes and plant roots via exudate secretion are not well understood (32).
Many studies on the rhizosphere microbiome have been conducted over the years. However, only a few of them have focused on the archaeal microbiome of roots (33–37). Although archaea are widespread in terrestrial environments (38–41), little is known about the selection forces that shape their composition as well as their functions, survival, and proliferation strategies in the rhizosphere (42). Nitrososphaerota (formerly known as Thaumarchaeota) are the predominant archaea found in soil (40, 43). Members of Nitrososphaerota belonging to groups I.1a, I.1a-associated, and I.1b (44) are ammonia-oxidizing archaea (AOA) involved in autotrophic ammonia oxidation, a key step in the nitrification process (43). Nitrification changes the availability of nitrogen species to plants and thus affects nitrogen fertilizer efficiency and enhances nitrogen mobility in the environment, resulting in fertilizer loss and eutrophication of water bodies. In addition, the nitrification intermediate, nitric oxide, functions as a signaling molecule in plants (45), and ammonia-oxidizing microorganisms (AOM) produce and emit nitrous oxide from agricultural soil (46).
Here, we analyzed archaeal communities associated with the rhizosphere of pepper and ginseng plants. The majority of the archaea identified in bulk and rhizosphere soils were AOA related, and they were found to outnumber ammonia-oxidizing bacteria (AOB) in both bulk and rhizosphere soils. Furthermore, AOA communities differed between bulk and rhizosphere soils, with the latter dominated by AOA closely related to “Candidatus Nitrosocosmicus,” a known manganese catalase (MnKat)-containing AOA. Based on these findings, we propose that “Candidatus Nitrosocosmicus” may be an important AOM associated with the rhizosphere.
RESULTS
16S rRNA gene amplicon-based archaeal community analysis in pepper and ginseng rhizosphere soils
The prokaryotic communities in bulk and rhizosphere soils of pepper plants were examined using 16S rRNA gene amplicon sequencing during the plant vegetative (60-day-old) and reproductive (90-day-old) growth phases. The compositions and diversities of prokaryotic communities in pepper plant rhizosphere soils differed from those in bulk soils regardless of growth phase (see more details in Supplementary Results and Discussion) (Fig. S1). Archaea were less abundant in rhizosphere soils (0.66 ± 0.23%; n = 10) compared to bulk soils (3.46 ± 0.64%; n = 15) in both plant growth phases (Table S1). Members of the phylum Nitrososphaerota, which were abundant in both the rhizosphere and bulk soils, accounted for a significant proportion of total archaeal 16S rRNA gene reads (94.2%; n = 25). They were also the 11th most abundant prokaryotic phyla detected (Fig. S1A; Table S1), with the majority of their 16S rRNA gene amplicon sequencing variants (ASVs) belonging to three AOA groups (I.1a, I.1a-associated, and I.1b). When compared to other AOM, such as Nitrosomonadaceae (the family that includes AOB) and Nitrospiraceae [the family that includes both complete ammonia oxidizers (Comammox) and nitrite oxidizers], ASVs belonging to AOA were the most abundant in both bulk and rhizosphere soils of pepper plants (Fig. S2A).
Interestingly, distinct AOA communities were observed on the non-metric multidimensional scaling (NMDS) plot between bulk and rhizosphere soils (Fig. 1A), with a low-stress value (Stress = 0.032), and ANOSIM analysis supported this difference (R = 0.997, P < 0.001). The difference between plant growth phases had no significant effect on the AOA communities in the rhizosphere soils (R = 0.033, P > 0.05). To identify ASVs that contributed to the discrimination of the AOA communities between bulk and rhizosphere soils, an indicator species analysis was performed at the ASV level. Among the AOA 16S rRNA gene ASVs, only 16S_ASV99 was significantly more abundant in the rhizosphere soils (IndVal: 0.79, q < 0.01) (Fig. 1B). This ASV is closely related to members of the clade “Ca. Nitrosocosmicus” in group I.1b (Fig. 2A), with >99.5% 16S rRNA sequence similarity. A comparison of the relative abundances of the AOA 16S rRNA gene ASVs showed that 16S_ASV99 was the single most abundant AOA ASV in the rhizosphere soils (an average of 44.4% of the total AOA 16S rRNA gene reads) (Fig. 1B). On the other hand, two AOA 16S rRNA gene ASVs were dominant in the bulk soils (16S_ASV17 and 16S_ASV1, accounting for 29.4% and 8.8%, respectively, of the total AOA 16S rRNA gene reads) (Fig. 1B), and they were closely related to fosmid clone 54d9 of group I.1b and the clade “Ca. Nitrosotenuis” of group I.1a, respectively (Fig. 2A).
Fig 1
Fig 2
For comparison, prokaryotic communities in bulk and rhizosphere soils of perennial ginseng plants (2-, 4-, and 6-year-old), collected from a different geographical region than the pepper plants, were analyzed (Table S2). As observed for pepper plants (Fig. S1A; Table S1), AOA were the most dominant archaea in both bulk and rhizosphere soils (Table S2). Rhizosphere AOA communities of the ginseng plants clustered based on the AOA 16S rRNA gene profiles (Fig. 1C) but were distinct from those of the bulk soils (Stress: 0.056; ANOSIM, R = 0.908, P < 0.001). The plant age had no significant effect on the AOA communities in the rhizosphere soils (ANOSIM, R = 0.158, P < 0.05). Notably, 16S_ASV38 related to “Ca. Nitrosocosmicus” was the most abundant ASV (accounting for 48.2% of the total AOA 16S rRNA gene reads) in the rhizosphere of the ginseng plants and strongly associated with rhizosphere soils than bulk soils (IndVal: 0.82, q < 0.01) (Fig. 1D and 2A) as observed in the pepper plants. In contrast to the pepper plants, two AOA ASVs were dominant in the bulk soils (16S_ASV233 and 16S_ASV741, accounting for 37.7% and 12.9%, respectively, of the total AOA 16S rRNA gene reads (Fig. 1D), and were closely related to the clade “Ca. Nitrosotalea” of group I.1a-associated (Fig. 2A).
AOA amoA gene amplicon-based AOA community analysis
The AOA amoA gene amplicon sequencing analysis supported the presence of distinct AOA communities in the rhizosphere vs bulk soils of pepper plants (Stress: 0.041; ANOSIM: R = 0.810, P < 0.001), while no significant difference was observed based on plant age (ANOSIM: R = −0.01, P > 0.05) (Fig. 3A). The number of amoA gene reads was summed for each AOA clade and compared between bulk and rhizosphere soils (Fig. 3B). Two “Ca. Nitrosocosmicus”-related amoA ASV clusters (amoA_cluster11, and −13) of clade NS-ζ (Zeta) (49) (Fig. 2B; Data set S1) had the highest relative abundance (average 39.7%; n = 10) (Fig. 3B) and strongly contributed to the clustering of the AOA communities in rhizosphere soils (Fig. 3A). On the other hand, the two most abundant bulk soil-associated AOA clade were NS-δ (Delta)−1 and NP-η (Eta)−1, respectively (Fig. 3B; Data set S1). These amoA clades correspond to the AOA clades fosmid clone 54d9 and “Ca. Nitrosotenuis,” respectively (Fig. 2B). These amoA-based results are highly consistent with the AOA 16S rRNA gene profiles (Fig. 1A and B).
Fig 3
In ginseng plants, the AOA amoA gene amplicon sequencing analysis also confirmed the clear segregation of the AOA communities between bulk and rhizosphere soils (Stress: 0.075; ANOSIM, R = 0.941, P < 0.001), as observed in pepper plants (Fig. 3A). However, plant age showed no significant effect on AOA communities (ANOSIM, R = 0.233, P < 0.01) (Fig. 3C). The relative abundance of the clade NS-ζ (amoA_ASV8, −34, and amoA_cluster12) significantly increased in the rhizosphere soils, while that of the clade NT-α−1 (i.e., I.1a-associated; amoA_cluster9) decreased (Fig. 3D; Data set S1), which was consistent with the results of the AOA 16S rRNA gene analysis (Fig. 1D). However, the notable increase in the relative abundance of the clades NS-δ−1 (amoA_cluster15, −16, and −17) and NP-η−1 (amoA_cluster5, −7 and −8) in the ginseng rhizosphere soils (Fig. 3D; Data set S1) was unexpected considering the AOA 16S rRNA gene analysis results (Fig. 1D).
Abundance of AOA MnKat gene in rhizosphere soils
Members of the genus “Ca. Nitrosocosmicus” possess genomic, morphological, and physiological properties distinct from other AOA (50–53). They possess genes that encode a putative MnKat (50–53) (Fig. S3). Despite the presence of MnKat genes in their genomes, catalase activity in these AOA has not yet been directly demonstrated experimentally. Nonetheless, the ability of “Ca. Nitrosocosmicus oleophilus” to grow in the absence of H2O2 scavengers, unlike other AOA, has already highlighted the presence of an active MnKat (50, 54). To test for catalase activity, we performed whole-cell assays with different AOMs. As expected, the heme-catalase-containing AOB strain (Nitrosomonas europaea ATCC 19718) (55), which was used as a positive control, generated O2 in the presence of H2O2 (Fig. 4A). Similarly, O2 generation in the presence of H2O2 was detected for “Ca. Nitrosocosmicus oleophilus” MY3 (Fig. 4B), consistent with the proposed activity of its MnKat. By contrast, O2 generation was not observed in the tested catalase gene-negative AOA, Nitrosarchaeum koreense MY1 (clade NP-γ (Gamma)−2.2; group I.1a), and Nitrososphaera viennensis EN76 (clade NS-α (Alpha)−3; group I.1b) (Fig. 4C and D). Based on these observations, we hypothesized that catalase activity can be one of the important factors associated with the dominance and survival of the “Ca. Nitrosocosmicus”-related AOA in the rhizosphere AOA community of pepper and ginseng plants. Coincidently, catalase-containing bacterial 16S rRNA gene ASVs were abundant in the rhizosphere soils of the pepper plants compared to bulk soils (Fig. S4; Data set S2 and S3). Specifically, 68.5% of the top 73 rhizosphere soil-associated ASVs (indicated by a positive value in log2fold) are known to have catalase genes or activity. By contrast, only 22.4% of the top 58 bulk soil-associated ASVs (indicated by a negative value in log2fold) have a catalase gene or activity (Fig. S4; Data set S2 and S3).
Fig 4
Thus, using qPCR, we quantified the relative abundance of MnKat-containing AOA in the rhizosphere soils of pepper and ginseng plants by comparing the copy numbers of the AOA MnKat genes relative to the amoA genes. Based on the newly designed PCR primer pair targeting AOA MnKat genes (see details in Materials and Methods), we found that all amplified MnKat gene sequences belonged to AOA, and more importantly, sequences associated with “Ca. Nitrosocosmicus”-related AOA were predominant in the rhizosphere and bulk soils (Table S3; Data set S4). In addition, AOA amoA gene copy numbers per gram of soil were lower in rhizosphere soils compared to bulk soils of the pepper plants (Fig. 5A). On the other hand, AOA MnKat gene copy numbers in 60- and 90-day-old rhizosphere soils were comparable to those in bulk soils (Fig. 5A). Overall, the copy number ratios of AOA MnKat genes to amoA genes were significantly higher in pepper plant rhizosphere soils than in bulk soils (Fig. 5B). In ginseng plants, the AOA amoA gene copy numbers per gram of soil were lower in rhizosphere soils compared to bulk soils only in 6-year-old plants, with similar AOA MnKat gene copy numbers in both soil types (Fig. 5C). In 4-year-old ginseng plants, AOA MnKat gene copy numbers were higher in rhizosphere soils than in bulk soils (Fig. 5C). The copy number ratios of AOA MnKat genes to amoA genes were consistently higher in rhizosphere soils than in bulk soils across all plant ages of ginseng plants (Fig. 5D). The results of AOA MnKat gene sequencing showed that, while sequences associated with the “Ca. N. oleophilus” MY3-like clade accounted for approximately 66%, and sequences associated with “Ca. N. everglandensis” SR1-like made up most of the remaining portion in bulk soil, in rhizosphere soils, the proportion of sequences associated with the “Ca. N. oleophilus” MY3-like clade increased to nearly 99% (Table S3). Overall results from both plants revealed that (i) most of the amplified MnKat gene sequences are associated with “Ca. Nitrosocosmicus” and (ii) the copy number ratios AOA MnKat genes to amoA genes were higher in the rhizosphere soils than in bulk soils.
Fig 5
Enrichment of MnKat-containing AOA from soil by H2O2 treatment
To investigate the selective enrichment of MnKat-containing AOA in H2O2-amended soils, bulk soil slurries from the pepper plant experiment were incubated with 1.5 mM ammonium chloride in the presence of 0–30 µM H2O2 (Fig. S5). Because H2O2 is rapidly decomposed by soil microorganisms and abiotic processes (Fig. S6), resulting in a short half-life (14.9 min at 10 µM H2O2), it was added to the soil slurries twice daily during the incubation period. Ammonia oxidation was gradually inhibited as the concentration of amended H2O2 in the soil slurry increased (Fig. S5). The copy number ratios of AOA MnKat genes to amoA genes increased in proportion to the concentration of H2O2 amended (Fig. S7), indicating that the presence of H2O2 in the soil selectively enriched catalase-containing AOA.
AOA community analysis of the soil slurries amended with H2O2 was performed using amplicon sequencing of prokaryotic 16S rRNA and AOA amoA genes (Fig. 6). We found a clear segregation between the slurry samples with different H2O2 concentrations based on the AOA 16S rRNA gene profiles (Stress: 0.023; ANOSIM, R = 0.822, P < 0.001). A cluster of the soil slurries with 30 µM H2O2 was especially distinct from the others (Fig. 6A). The relative abundances of 16S_ASV38 and −99 belonging to the clade “Ca. Nitrosocosmicus” (Fig. 3A) increased proportionally to the H2O2 concentration based on the AOA 16S rRNA gene amplicon sequencing results (Fig. 6B). By contrast, the relative abundance of other AOA 16S rRNA gene ASVs (16S_ASV1, -2, and -58), belonging to the clades Nitrosarchaeum and “Ca. Nitrosotenuis” of group I.1a (Fig. 3A), significantly decreased as the concentration of H2O2 increased (Fig. 6B). The relative abundance of 16S_ASV17, affiliated with the clade fosmid clone 54d9, remained unchanged across all H2O2 concentrations (Fig. 6B).
Fig 6
The segregation of AOA communities in the soil slurry samples with varying H2O2 concentrations was also supported by AOA amoA gene amplicon sequencing data (Stress: 0.067; ANOSIM, R = 0.649, P < 0.001) (Fig. 6C). The soil slurry sample cluster with 30 µM H2O2 was especially distinct from the others (Fig. 6C); this finding was also consistent with the AOA 16S rRNA gene-based analysis (Fig. 6A). The relative abundances of amoA clades NS-δ−1, NS-ζ, NS-α−3.2, and NS-δ−2 increased as H2O2 concentrations increased (Fig. 6D). Both the AOA 16S rRNA and amoA gene profiles revealed a significant increase in the relative abundance of “Ca. Nitrosocosmicus”-related AOA (clade NS-ζ) (Fig. 6B and D). Surprisingly, as the H2O2 concentrations increased, so did the relative abundance of clade NS-δ−1 amoA genes (related to the amoA gene on the fosmid clone 54d9), contrary to what was observed in the AOA 16S rRNA gene profile analysis (Fig. 6B). On the other hand, the relative abundance of other AOA clades in group I.1a (amoA clades NP-η-1 and NP-γ-2.2) decreased with increasing H2O2 concentrations (Fig. S6D), as observed in the AOA 16S rRNA gene profile analysis (Fig. 3A; Fig. 6B). Taken together, these results support the hypothesis that resistance to H2O2 is important in the selection of MnKat-containing AOA in soil habitats with elevated H2O2 concentrations, such as the plant rhizosphere.
Expression of AOA MnKat gene in rhizosphere soils
To demonstrate AOA MnKat gene expression in rhizosphere soils, transcripts of AOA MnKat genes were quantified using cDNA generated from mRNA extracted from pepper plant rhizosphere soils and bulk soils. In addition, a “Ca. Nitrosocosmicus” clade-specific housekeeping gene, rpoB, and AOA amoA gene were quantified to estimate the relative abundance of AOA MnKat gene transcripts in rhizosphere soils. The RT-qPCR data revealed that the relative abundance of AOA MnKat gene transcripts to AOA amoA and “Ca. Nitrosocosmicus” clade rpoB gene transcripts in rhizosphere soils was not significantly different from those in bulk soils (Fig. S8), indicating possible constitutive expression of AOA MnKat genes in soils.
DISCUSSION
There have been extensive studies on rhizosphere microbial communities, as the rhizosphere microbiome affects the survival of plants under stress conditions such as those caused by climate change, pathogen infection, etc. (1–3). Despite their potential importance in plant growth and development, archaea are only rarely included in rhizosphere microbiome studies (33–37). AOA are especially abundant in terrestrial environments and play a key role in the soil nitrogen cycle (33, 40, 43), necessitating additional research into their interactions with plant roots (33, 40). Patterns of prokaryotic communities formed in the analyzed rhizosphere soils (see more details in Supplementary Results and Discussion) (Fig. S1; Tables S1 and S2) distinct from the bulk soils of pepper plants were consistent with previous studies on other plant species (4, 8, 9, 56). Furthermore, distinct AOA communities in rhizosphere soils of pepper and ginseng plants relative to bulk soils were revealed by 16S rRNA and amoA genes amplicon sequencing profiles (Fig. 1A, C, 2A and C), indicating a niche differentiation of AOA between bulk and rhizosphere soils of the plants.
We observed a decrease in the relative abundance of AOA in rhizosphere soils of pepper and ginseng plants compared to bulk soils (Fig. S1A; Tables S1 and S2), which contradicts previous findings (57, 58). Although AOA had a low relative abundance in rhizosphere soils of pepper and ginseng plants, they still outnumbered other AOM (Fig. S2). Despite extensive studies on ecology of AOM in various soils, only limited studies were conducted for ecological factors affecting niche differentiation of clades of AOM in rhizosphere soils. It was known that traits of plants and N-fertilization can affect the relative abundance of AOA to AOB in the rhizospheres of the plants compared to AOB (58–60). Further research on biotic and abiotic rhizosphere factors affecting the differential abundance of AOA and AOB is warranted. Consequently, a low abundance of AOM may decrease nitrification activity near plant roots, which is desirable to reduce N losses and increase N fertilizer use efficiency (59, 61).
AOA related to “Ca. Nitrosocosmicus” were notably the most abundant in the rhizosphere soils based on 16S rRNA and amoA gene amplicon analyses (Fig. 1B, D, 2, 3B and 3D; Data set S1). Interestingly, among genome-sequenced AOA, MnKat genes are exclusively present in members of the genus “Ca. Nitrosocosmicus” and of the species “Ca. Nitrososphaera evergladensis” (50–52, 62) (Fig. 2). Phylogenetic analysis of MnKat genes (Fig. S3) revealed that Nitrososphaerota MnKat genes were closely related to those found in the bacterial phylum Terrabacteria, which includes common soil bacteria such as Actinomycetota and Bacillota (53) (Fig. S3). In addition, these genes differed from those found in closely related archaeal phyla, Ca. Thermoproteota and Euryarchaeota, implying that horizontal gene transfer events between archaea and bacteria shaped the evolutionary history of the MnKat gene (Fig. S3).
Catalase activity was measured in “Ca. Nitrosocosmicus oleophilus” MY3 (Fig. 4B), a strain closely related to AOA that was dominant in pepper and ginseng rhizospheres (Fig. 1B, D, 2, 3B and 3D; Data set S1). The AOA MnKat, whose active site is predicted to be stable under low H2O2 levels compared with the heme catalase (63), may provide an evolutionary advantage at low H2O2 levels (< 3 µM), which can completely inhibit the nitrification activity of catalase-negative AOA (64, 65). Based on the documented selection of MnKat-encoding AOA in rhizospheres of pepper and ginseng plants, as well as the experimental confirmation of catalase activity in a related AOA isolate, it is tempting to speculate that resistance to H2O2 is one of the important factors shaping AOA communities in rhizospheres. Consistently, we observed that the copy number ratios of the AOA MnKat gene to the amoA gene were significantly higher in rhizosphere soils of pepper and ginseng plants than in bulk soils (Fig. 5B and D). The dominance of AOA MnKat gene sequences closely related to “Ca. Nitrosocosmicus” in rhizosphere soils (Table S3) corroborated the results of AOA 16S rRNA and amoA gene analyses (Fig. 1 and 3).
Soil characteristics (4–9) and host phylogeny (4, 13, 21, 22, 24) are considered important determinants of rhizosphere microbial community composition and function. Even plant genotype-specific microbial communities have been observed in the rhizosphere of some plant species (5, 7). Despite the different life cycles, phylogeny, and geographic locations of the pepper and ginseng plants studied here, distinct AOA communities in the rhizosphere soils relative to the bulk soils were observed, which was attributable to the predominance of the MnKat-containing members of “Ca. Nitrosocosmicus” (Fig. 1B, D, 2, 3B and 3D; Data set S1). In this context, it is important to note that the dominant phylotype, C1b.A1 (representing the clones TRC23-30 and TRC23-38), belonging to Nitrososphaerota (formerly known as Crenarchaeota), was found to predominantly colonize the roots of tomato (Solanum lycopersicum L. in the order Solanales) grown in soil from a Wisconsin field (66). This phylotype is closely related to “Ca. Nitrosocosmicus oleophilus” MY3 with 99.7% 16S rRNA gene sequence similarity, suggesting that closely related members of “Ca. Nitrosocosmicus” are dominant in various agriculturally important plants and that the enrichment of the AOA in the plant rhizosphere may be widespread, regardless of geographical location and plant phylogeny.
In addition to the amoA clade NS-ζ containing members of the genus “Ca. Nitrosocosmicus,” the AOA amoA gene reads of the clade NS-δ-1 harboring the fosmid clone 54D9 amoA sequence were also abundant in the H2O2-amended soil slurries (Fig. 6D) and the rhizosphere soils of ginseng plants (Fig. 3D), but not in the pepper plants (Fig. 3B). It is yet unknown whether clade NS-δ-1 members have MnKat genes. The prominent increases in the relative abundance of the AOA amoA gene reads from clade 54D9 (Fig. 3D and 6D) are in stark contrast with the findings from the analysis of AOA 16S rRNA gene amplicon reads (Fig. 1D and 2A; Fig. 6B). Thus, we cannot rule out the possibility that the PCR primer set used to construct the AOA amoA gene amplicon libraries is biased towards clade NS-δ-1 amoA genes.
Oxygen supply is crucial for plant roots, not only for cell respiration but also for the formation of reactive oxygen species, including H2O2. H2O2 is a ubiquitous metabolic by-product of aerobic unicellular and multicellular organisms (67–70) that plays an important role in developmental and physiological processes in plant roots. H2O2 is involved in loosening cell walls for cell elongation in roots via peroxidase-mediated lignin formation (71, 72) and accumulates in the differentiation zone and the cell wall of root hairs during the formation of fine roots in Arabidopsis (Arabidopsis thaliana in the order Brassicales) (73). It was observed that H2O2 production increased in a specific region of fine roots after K+ deprivation (74). Similarly, H2O2 release from seedling roots into the environment has been observed (73, 75–77). Furthermore, mycorrhizae mediated an increase in H2O2 release from the roots of trifoliate orange to alleviate drought stress (78). Recently, it was proposed that the rhizosphere is a widespread but previously unappreciated hotspot for ROS production, with hydroxyl radicals, which represent ROS species, periodically accumulating up to >2 µM in rice plant rhizosphere soil pore water after 6 h of light exposure (79). Thus, plant roots trigger the release of H2O2 into their surroundings and thereby chemically shape the rhizosphere habitat. In addition to plant roots, soil microorganisms are known to release ROS (80, 81).
In the soil slurry experiments, we demonstrated that H2O2 amendment in bulk soils increased the abundance of MnKat-containing AOA in a concentration-dependent manner (Fig. S7). In addition, AOA from the clade “Ca. Nitrosocosmicus” became dominant in H2O2-amended soil slurries (Fig. 6B), and the copy number ratios of AOA MnKat genes to amoA genes increased as the concentration of H2O2 increased (Fig. S7B). The toxic effects of H2O2 on AOA were previously assessed with group I.1a, where ammonia oxidation was completely inhibited at levels of 0.2–3.0 µM H2O2 (64, 65). Consistently, the nitrification activity and abundance of AOA decreased in H2O2-amended soil slurries (Fig. S5 and S6). This might explain the decrease in gross nitrification rates in the rhizosphere (82). Furthermore, “Ca. Nitrosocosmicus” MnKat genes were found to be constitutively expressed in pepper plant rhizosphere soils and bulk soils (Fig. S8). Taken together, our results imply that rhizosphere H2O2 may be an important factor in the selection of MnKat-containing AOA in the plant rhizosphere. Interestingly, metagenomic and metatranscriptomic analyses of the rhizosphere microbial communities of cucumber (Cucumis sativus L. in the order Cucurbitales) and wheat (Triticum turgidum L. in the order Poales) plants identified the enrichment and expression of prokaryotic catalase genes, which were suggested to be associated with root colonization (83). The dominance of catalase-containing bacterial ASVs in rhizosphere soils of pepper plants over bulk soils (Fig. S4; Data set S2 and S3) observed in this study corresponds to these findings. It is plausible that H2O2 levels in rhizosphere environments may be inhibitory to catalase-negative microbes such as group I.1a and I.1a-associated AOA. In suspended aquatic environments, the growth of catalase-negative AOA could be supported by coexisting catalase-positive microbes (65, 84, 85). Hence, further study will be needed to reveal if such an interaction exists between catalase-positive microbes and catalase-negative AOA in soil environments. Therefore, we propose that the catalase activity of microorganisms in rhizospheres may serve as a microbial stress response. It may also modulate developmental and physiological processes in plant roots, as well as redox dynamics and biogeochemical processes in soil.
Despite the presence of the MnKat gene in the “Ca. Nitrososphaera evergladensis” genome, ASVs related to this AOA were not dominant in the analyzed microbial community in rhizosphere soils (Table S3). Thus, while catalase activity is a very plausible explanation for the selection of members of the genus “Ca. Nitrosocosmicus” in the rhizosphere, it should be noted that the genomes of these ammonia-oxidizers also encode various distinct traits that may individually or collectively confer higher rhizosphere fitness compared to other AOA. For example, tolerance to high salinity (51) and acidic pH (86), as well as the ability for biofilm formation (50) observed in “Ca. Nitrosocosmicus” members, may support survival in the rhizosphere and/or help establish interactions with plant roots. Furthermore, the higher concentration of ammonia in rhizosphere soils compared to bulk soils (87) might facilitate the competitive success of members of “Ca. Nitrosocosmicus,” which possess a lower affinity and lower specific affinity for ammonia than other AOA (88) in the rhizosphere. Thus, more research is needed to determine how catalase activity contributes to the enrichment of “Ca. Nitrosocosmicus” members in the rhizospheres of various plants.
The nitrification process, which converts ammonia to nitrite and then to nitrate, strongly affects the availability of nitrogen species for plant roots (89). The available inorganic nitrogen species ratio (ammonium:nitrate) is significant to plant growth by influencing cellular pH maintenance and energy efficiency of nitrogen assimilation in plants (90, 91). Due to their abundance, AOA, especially catalase-containing “Ca. Nitrosocosmicus” members, as demonstrated in this study, are considered to be important players mediating the nitrification process in the rhizosphere (59). Song et al. demonstrated that “Ca. Nitrosocosmicus oleophilus” MY3 cells colonized the root surface of Arabidopsis plants and proposed that volatile compounds emitted by “Ca. Nitrosocosmicus oleophilus” MY3 could elicit induced systemic resistance (92). Taken together, the selection of catalase-containing AOA of the genus “Ca. Nitrosocosmicus” in the rhizosphere of several agriculturally important plants hints at a previously overlooked AOA-plant interaction. Our understanding of AOA-plant interactions in the rhizosphere is still in its infancy, and this study highlights an important clade of AOA with already available cultured representatives for further mechanistic analyses in this crucial research field.
MATERIALS AND METHODS
Plant cultivation, soil collection, and analysis
Soil samples from the bulk and rhizosphere of pepper (Capsicum annuum L., Solanaceae) and ginseng (Panax ginseng C.A. Meyer, Araliaceae) plants were collected to investigate the plant-root-associated prokaryotic communities. The pepper plants were grown in sandy loam soil under a rain shelter for 6 months (March to September 2017), with the addition of ammonium sulfate (55 kg N ha−1) as a nitrogen fertilizer once before transplanting. After transplanting, the average temperature in the rain shelter was kept at 25 ± 5°C, and the soil was watered daily as needed to keep soil moisture above 20% at 30 cm soil depth. A total of 25 soil samples were collected: bulk soil (n = 5) at day 0, and both bulk and rhizosphere soils (n = 5 each) at 60 days (vegetative phase) and 90 days (reproductive phase). The rhizosphere soil samples were collected as soil tightly adhering to plant roots. Five bulk soil subsamples were collected at 15 cm soil depth and 40 cm away from the plants and then mixed. The soil samples were stored at −80°C until DNA extraction. The locations and general properties of bulk soil are presented in Table S4.
The ginseng plants were grown for 6 years (2012–2018) after transplanting 1-year-old seedlings (Table S4) to a sandy loam soil field. The Ginseng Good Agricultural Practices Scheme from the National Institute of Horticultural and Herbal Science of the Rural Development Administration (Republic of Korea) was followed for pre-planting treatment, pest management, watering, and fertilization. During the cultivation period, a total of 29 soil samples were collected: bulk soils (n = 3 for each age group) and rhizosphere soils (n = 4, 7, and 9 for 2-, 4-, and 6-year-old ginseng plants, respectively). The collection of soil samples and analysis of general properties of soils were conducted in the same manner as described above for pepper plants.
DNA extraction and quantification of AOA amoA and MnKat genes
Genomic DNA was extracted from each 0.25 g soil sample using the Exgene Soil DNA mini extraction kit (GeneAll Biotechnology Co. Ltd., Republic of Korea) according to the manufacturer’s instructions. The DNA concentration and purity were measured using a Nanodrop ND-1000 spectrophotometer (NanoDrop Technologies, USA). To check for possible quantitative PCR (qPCR) or PCR inhibition, genomic DNA from Methylacidiphilum caldifontis IT6 (106 gene copy numbers per reaction) (93) was spiked as an internal positive control (94) into serially diluted template DNA (0.25–20 ng) extracted from soils of the pepper and ginseng plants. Strain IT6-specific pmoA1 gene primer pair was used for qPCR detection. The quantitative results of the spiked control within the template or pure water were assessed by comparing the Ct values of the pmoA1 gene. No significant PCR inhibition was observed in the serially diluted template DNA (0.25–20 ng), and thus, we used<10 ng DNA for further qPCR analysis.
The copy numbers of the AOA amoA gene were assessed using the CrenamoA104F/CrenamoA616R primer pairs (see Table S5). The primer pair for quantifying the AOA MnKat gene (aoa-MnKat200F: 5′-GAAGAGATRGGWCATGTWGA-3′, aoa-MnKat480R: 5′-CCTGTMGCYTCAAGCATDA-3′) was newly designed using MnKat gene sequences retrieved from the genomes of “Ca. Nitrosocosmicus oleophilus” MY3 (GCA_000802205.2), “Ca. Nitrosocosmicus arcticus” Kfb (GCA_007826885.1), “Ca. Nitrosocosmicus hydrocola” G61 (GCF_001870125.1), and “Ca. Nitrososphaera evergladensis” SR1 (GCA_000730285.1). 1–10 ng of sample DNA was used for qPCR using a MiniOpticon qPCR detection system (Bio-Rad Laboratories, Hercules, USA). The iQ SYBR Green Supermix (Bio-Rad Laboratories, USA) and PCR primer pairs were used for PCR amplification. AOA amoA gene was amplified via the following steps: 95°C for 3 min; followed by 40 cycles at 95°C for 45 s, 55°C for 45 s, 72°C for 45 s; and 72°C for 5 min. AOA MnKat gene was amplified via the following steps: 95°C for 3 min; followed by 40 cycles at 95°C for 45 s, 55°C for 45 s, 72°C for 45 s; and 72°C for 5 min. A dilution series of genomic DNA from strain MY3 was included in every qPCR cycle for calibration purposes. Amplification efficiencies ranged between 80% and 94% for all target genes, and qPCR R2 calibration values were greater than 0.99.
Amplicon sequencing and phylogenetic analysis
For the construction of amplicon sequencing libraries, the 16S rRNA gene was amplified using the 515F/926R primer pairs (see Table S6), via the following steps: 95°C for 3 min; followed by 25 cycles at 95°C for 45 s, 50°C for 45 s, 72°C for 90 s; and 72°C for 5 min. AOA amoA gene was amplified with the CrenamoA104F/CrenamoA616R primer pairs (see Table S5), via the following steps: 95°C for 3 min; followed by 30 cycles at 95°C for 45 s, 55°C for 45 s, 72°C for 45 s; and 72°C for 5 min. AOA MnKat gene was amplified with the aoa-MnKat200F/aoa-MnKat480R primer pairs via the following steps: 3 min heating step at 95°C; followed by 35 cycles at 95°C for 45 s, 55°C for 45 s, 72°C for 45 s; and 72°C for 5 min. The following index PCR for both gene libraries was conducted with the Nextera XT index kit v2 (Illumina Inc., USA). The PCR product was purified using the Labopass DNA purification kit (Cosmogenetech Inc., Republic of Korea). The sequencing was performed using the Illumina MiSeq (2 × 300 bp) platform (Illumina Inc., USA) at Macrogen Inc. (Republic of Korea). The QIIME2 (v2024.2) pipeline with implemented tools for quality control (Cutadapt) (95) and de-noising, and pair read merging (DADA2) (96) was used to analyze the amplicon sequence data. The primer region was trimmed. After quality plots were generated, the sequences failing to pass an average base call accuracy of 99% (median Phred score of 20) were excluded. Low-quality regions of each sequence were removed during the de-noising step using DADA2 with the following parameters: 16S rRNA gene: --p-trunc-len-f 264 --p-trunc-len-r 168 --p-max-ee-f 2 --p-max-ee-r 4; AOA amoA gene: --p-trunc-len-f 280 --p-trunc-len-r 265 --p-max-ee-f 2 --p-max-ee-r 4; AOA MnKat gene: --p-trunc-len-f 188 --p-trunc-len-r 110 --p-max-ee-f 3 --p-max-ee-r 3(94). ASVs were taxonomically classified using qiime feature-classifer and sklearn algorithm against Greengene2 database (v2022.10) for the 16S rRNA gene, the Alves RJE et al. (49) reference data set for the AOA amoA gene, and AOA MnKat genes. For further analyses, the 16S rRNA and the AOA amoA gene ASV tables were rarefied to even depths of 42,584 and 43,680 reads, respectively.
For phylogenetic analyses, representative full-length sequences of the AOA 16S rRNA, amoA, and MnKat genes were obtained from the NCBI database. Alignments of the derived sequences were performed using MAFFT (v7.453) (97). A maximum likelihood phylogenetic tree was constructed with IQ-TREE (v1.6.12) (48).
Catalase activity assays
Three strains of AOA (“Ca. Nitrosocosmicus oleophilus” MY3, Nitrosoarchaeum koreense MY1, and Nitrososphaera viennensis EN76) and one strain of AOB (Nitrosomonas europaea ATCC 19718) were grown under optimal conditions (98). After oxidizing 1 mM ammonia, the cells were harvested by centrifugation (5,000 × g for 20 min at 25°C). The harvested cells were washed three times using a basal artificial freshwater medium (AFM) (98), which is devoid of ammonia, trace elements, and pyruvate. Subsequently, 2 mL aliquots of cell suspension were filled into a respiration chamber fitted with contactless oxygen sensor spots (OXSP5, PyroScience, Germany) and maintained for 10–20 min before H2O2 injection. The FireSting fiber-optical oxygen meter FSO2-1 (PyroScience, Germany) operation and two‐point calibration followed the manufacturer’s instructions. The reaction chambers were set at the optimal growth temperature for each strain (98) in a water bath (NB-304, N-BIOTEK, Republic of Korea) and stirred with a magnetic stirrer (MIXdrive 1 XS, 2mag AG, Germany) at 1,000 rpm.
Soil slurry experiment
For soil slurry experiments, the bulk soil composited from five subsamples from the pepper plant experiment was sieved (2-mm mesh size) to remove plant debris and stones. Aliquots of the fresh bulk soil (0.25 g) were incubated in 25 cm3 cell culture flasks (SPL Life Sciences, Republic of Korea) containing 10 mL of AFM. The medium contained NH4Cl (1.5 mM), NaHCO3 (2 mM), and MES buffer (pH 6.5; 3 mM). Allylthiourea (50 µM), an inhibitor of AOB growth (99), was used to focus on the response of soil AOA to H2O2 amendment. The cultures were incubated under aerobic conditions in the dark at 30°C in a static incubator with intermittent mixing. During the experiment, H2O2 was amended twice daily to reach final concentrations of 0–30 µM. Nitrite and nitrate were measured using spectrophotometric methods as previously described (98) for the estimation of nitrification activity. For DNA extraction, soil aliquots were obtained from the slurry samples by centrifugation (8,000 × g for 20 min at 25°C) and were stored at −80°C.
To estimate the biotic and abiotic decomposition rates of H2O2 in soil slurries, H2O2 concentrations in fresh bulk soils and autoclaved soil slurries were measured using ISO-HPO-100 (WPI, UK) amperometric sensors, with an Apollo 4000 System (WPI, UK). For H2O2 measurements, soil slurries were stirred using an MS-500 magnetic stirrer (Duksan General Science Co., Republic of Korea) at 1,500 rpm. The electrode was calibrated against freshly prepared H2O2 solutions in the range of 0–10 µM in AFM at 25°C.
AOA MnKat gene expression assay
Total RNA was extracted from each 2 g of bulk and rhizosphere soil samples (bulk soil: n = 5, rhizosphere soil: n = 5, from 90-day-old pepper plants) using the RNeasy PowerSoil Total RNA Kit (Qiagen, USA) according to the manufacturer’s instructions. RNA was eluted in 100 µL of RNase-free water. After removing the DNA from the eluent by treatment with DNase, a SuperScript IV VILO ezDNAse kit (Thermo Fisher Scientific, USA) was used for cDNA synthesis. The concentrations of RNA and cDNA were determined using a Qubit 4 fluorometer (Thermo Fisher Scientific, USA).
The relative expression of the AOA MnKat gene in bulk and rhizosphere soils was estimated using a “Ca. Nitrosocosmicus” clade-specific housekeeping gene, rpoB, and AOA amoA gene (see Table S5). The primer pair for quantifying the rpoB gene (nsc-rpoB2884F: 5′-TAYGGWTTYAAGCAYAGTGG-3′, nsc-rpoB3320R 5′-TGAGTTTAAATGTSGCWCC-3′) was newly designed using rpoB sequences retrieved from “Ca. Nitrosocosmicus” clade-genomes (see Fig. S9). The “Ca. Nitrosocosmicus” clade-specific rpoB gene was amplified via the following steps: 95°C for 3 min; followed by 40 cycles at 95°C for 45 s, 59°C for 45 s, 72°C for 45 s; and 72°C for 5 min.
Statistical analysis
All statistical analyses were conducted using the R statistical software (v4.1.2) and R Studio (v2022.02.3). Microbial diversity analysis and data visualization were performed using the R packages phyloseq (v1.26.0) (100), vegan (v2.5–3) (101), and ggplot2 (v3.1.0) (102). Processed amplicon sequence reads were imported using phyloseq (100). NMDS analysis and principal coordinate analysis (PCoA) based on Bray–Curtis dissimilarity metrics were used in Vegan (101) to compare the microbial communities between the samples. The ordination analysis patterns were statistically tested using permutational multivariate analysis of variance (PERMANOVA) and analysis of similarity (ANOSIM) with adonis2 and vegdist, respectively, which are part of the vegan packages in R (101). Indicator species were identified using the indval function of labdsv package (v2.0–1) (103). Samples from the same compartment were treated as a group for indicator species analysis.
ACKNOWLEDGMENTS
This work was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Korean government (Ministry of Education) (2020R1A6A1A06046235), the NRF grants funded by the Korean government (Ministry of Science and ICT) (2021R1A2C3004015), the National Institute of Agricultural Science, Ministry of Rural Development Administration, Republic of Korea (PJ01700703), and the Korea Institute of Marine Science & Technology Promotion (KIMST) funded by the Ministry of Oceans and Fisheries (RS-2024-00436293). J.-H.G. was supported by the NRF grant funded by the MSIT (RS-2023–00213601). M.-Y.J. was supported by the NRF grant funded by MSIT (NRF-2021R1C1C1008303 and NRF-2022R1A4A503144711).
U.-J.L., J.-H.G., S.C., and S.-K.R. conceptualized the study. U.-J.L., J.-H.G., and S.C. performed the investigation. U.-J.L., J.-H.G., M.-Y.J., and S.-K.R. performed formal analysis and visualized the study. M.-Y.J., T.K.L., H.R., Z.-X.Q., and S.-K.R. provided comments on methodology and validation of the study. U.-J.L., J.-H.G., and S.-K.R. wrote the original draft. U.-J.L., J.-H.G., M.-Y.J., T.K.L., H.R., S.I.A., W.W., M.W., Z.-X.Q., and S.-K.R. reviewed and edited the manuscript.
SUPPLEMENTAL MATERIAL
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REFERENCES
1.
Lemanceau P, Barret M, Mazurier S, Mondy S, Pivato B, Fort T, Vacher C. 2017. Chapter 5, Plant communication with associated microbiota in the spermosphere, rhizosphere and phyllosphere, p 101–133. In Becard G (ed), Advances in botanical research, 1st ed. Academic Press.
2.
Korenblum E, Dong Y, Szymanski J, Panda S, Jozwiak A, Massalha H, Meir S, Rogachev I, Aharoni A. 2020. Rhizosphere microbiome mediates systemic root metabolite exudation by root-to-root signaling. Proc Natl Acad Sci U S A 117:3874–3883.
3.
Pascale A, Proietti S, Pantelides IS, Stringlis IA. 2019. Modulation of the root microbiome by plant molecules: the basis for targeted disease suppression and plant growth promotion. Front Plant Sci 10:1741.
4.
Bulgarelli D, Rott M, Schlaeppi K, Ver Loren van Themaat E, Ahmadinejad N, Assenza F, Rauf P, Huettel B, Reinhardt R, Schmelzer E, Peplies J, Gloeckner FO, Amann R, Eickhorst T, Schulze-Lefert P. 2012. Revealing structure and assembly cues for Arabidopsis root-inhabiting bacterial microbiota. Nat New Biol 488:91–95.
5.
Lundberg DS, Lebeis SL, Paredes SH, Yourstone S, Gehring J, Malfatti S, Tremblay J, Engelbrektson A, Kunin V, Del Rio TG, Edgar RC, Eickhorst T, Ley RE, Hugenholtz P, Tringe SG, Dangl JL. 2012. Defining the core Arabidopsis thaliana root microbiome. Nat New Biol 488:86–90.
6.
Peiffer JA, Spor A, Koren O, Jin Z, Tringe SG, Dangl JL, Buckler ES, Ley RE. 2013. Diversity and heritability of the maize rhizosphere microbiome under field conditions. Proc Natl Acad Sci U S A 110:6548–6553.
7.
Bulgarelli D, Garrido-Oter R, Münch PC, Weiman A, Dröge J, Pan Y, McHardy AC, Schulze-Lefert P. 2015. Structure and function of the bacterial root microbiota in wild and domesticated barley. Cell Host Microbe 17:392–403.
8.
Edwards J, Johnson C, Santos-Medellín C, Lurie E, Podishetty NK, Bhatnagar S, Eisen JA, Sundaresan V. 2015. Structure, variation, and assembly of the root-associated microbiomes of rice. Proc Natl Acad Sci USA 112:E911–E920.
9.
Schreiter S, Ding G-C, Heuer H, Neumann G, Sandmann M, Grosch R, Kropf S, Smalla K. 2014. Effect of the soil type on the microbiome in the rhizosphere of field-grown lettuce. Front Microbiol 5:144.
10.
Paterson E, Gebbing T, Abel C, Sim A, Telfer G. 2007. Rhizodeposition shapes rhizosphere microbial community structure in organic soil. New Phytol 173:600–610.
11.
Marschner H, Römheld V, Horst WJ, Martin P. 1986. Root‐induced changes in the rhizosphere: importance for the mineral nutrition of plants. J Plant Nutr Soil 149:441–456.
12.
Dennis PG, Miller AJ, Hirsch PR. 2010. Are root exudates more important than other sources of rhizodeposits in structuring rhizosphere bacterial communities? FEMS Microbiol Ecol 72:313–327.
13.
Haichar FEZ, Marol C, Berge O, Rangel-Castro JI, Prosser JI, Balesdent J, Heulin T, Achouak W. 2008. Plant host habitat and root exudates shape soil bacterial community structure. ISME J 2:1221–1230.
14.
Berendsen RL, Pieterse CMJ, Bakker PAHM. 2012. The rhizosphere microbiome and plant health. Trends Plant Sci 17:478–486.
15.
Hu L, Robert CAM, Cadot S, Zhang X, Ye M, Li B, Manzo D, Chervet N, Steinger T, van der Heijden MGA, Schlaeppi K, Erb M. 2018. Root exudate metabolites drive plant-soil feedbacks on growth and defense by shaping the rhizosphere microbiota. Nat Commun 9:2738.
16.
DeAngelis KM, Brodie EL, DeSantis TZ, Andersen GL, Lindow SE, Firestone MK. 2009. Selective progressive response of soil microbial community to wild oat roots. ISME J 3:168–178.
17.
Vranova V, Rejsek K, Skene KR, Janous D, Formanek P. 2013. Methods of collection of plant root exudates in relation to plant metabolism and purpose: a review. Z Pflanzenernähr Bodenk 176:175–199.
18.
Pausch J, Kuzyakov Y. 2018. Carbon input by roots into the soil: quantification of rhizodeposition from root to ecosystem scale. Glob Chang Biol 24:1–12.
19.
Pausch J, Tian J, Riederer M, Kuzyakov Y. 2013. Estimation of rhizodeposition at field scale: upscaling of a 14C labeling study. Plant Soil 364:273–285.
20.
Olanrewaju OS, Ayangbenro AS, Glick BR, Babalola OO. 2019. Plant health: feedback effect of root exudates-rhizobiome interactions. Appl Microbiol Biotechnol 103:1155–1166.
21.
Carvalhais LC, Dennis PG, Badri DV, Kidd BN, Vivanco JM, Schenk PM. 2015. Linking jasmonic acid signaling, root exudates, and rhizosphere microbiomes. Mol Plant Microbe Interact 28:1049–1058.
22.
Okubo A, Matsusaka M, Sugiyama S. 2016. Impacts of root symbiotic associations on interspecific variation in sugar exudation rates and rhizosphere microbial communities: a comparison among four plant families. Plant Soil 399:345–356.
23.
Pacheco-Moreno A, Bollmann-Giolai A, Chandra G, Brett P, Davies J, Thornton O, Poole P, Ramachandran V, Brown JKM, Nicholson P, Ridout C, DeVos S, Malone JG. 2024. The genotype of barley cultivars influences multiple aspects of their associated microbiota via differential root exudate secretion. PLoS Biol 22:e3002232.
24.
Li M, Song Z, Li Z, Qiao R, Zhang P, Ding C, Xie J, Chen Y, Guo H. 2022. Populus root exudates are associated with rhizosphere microbial communities and symbiotic patterns. Front Microbiol 13:1042944.
25.
Reinhold-Hurek B, Bünger W, Burbano CS, Sabale M, Hurek T. 2015. Roots shaping their microbiome: global hotspots for microbial activity. Annu Rev Phytopathol 53:403–424.
26.
Sasse J, Martinoia E, Northen T. 2018. Feed your friends: do plant exudates shape the root microbiome? Trends Plant Sci 23:25–41.
27.
Pieterse CMJ, Van der Does D, Zamioudis C, Leon-Reyes A, Van Wees SCM. 2012. Hormonal modulation of plant immunity. Annu Rev Cell Dev Biol 28:489–521.
28.
Piasecka A, Jedrzejczak-Rey N, Bednarek P. 2015. Secondary metabolites in plant innate immunity: conserved function of divergent chemicals. New Phytol 206:948–964.
29.
Bamji SF, Corbitt C. 2017. Glyceollins: soybean phytoalexins that exhibit a wide range of health-promoting effects. J Funct Foods 34:98–105.
30.
Sarr PS, Ando Y, Nakamura S, Deshpande S, Subbarao GV. 2020. Sorgoleone release from sorghum roots shapes the composition of nitrifying populations, total bacteria, and archaea and determines the level of nitrification. Biol Fertil Soils 56:145–166.
31.
Preston GM. 2017. Profiling the extended phenotype of plant pathogens. Mol Plant Pathol 18:443–456.
32.
Mendes R, Garbeva P, Raaijmakers JM. 2013. The rhizosphere microbiome: significance of plant beneficial, plant pathogenic, and human pathogenic microorganisms. FEMS Microbiol Rev 37:634–663.
33.
Moissl-Eichinger C, Pausan M, Taffner J, Berg G, Bang C, Schmitz RA. 2018. Archaea are interactive components of complex microbiomes. Trends Microbiol 26:70–85.
34.
Taffner J, Bergna A, Cernava T, Berg G. 2020. Tomato-associated archaea show a cultivar-specific rhizosphere effect but an unspecific transmission by seeds. Phytobiomes J 4:133–141.
35.
Karlsson AE, Johansson T, Bengtson P. 2012. Archaeal abundance in relation to root and fungal exudation rates. FEMS Microbiol Ecol 80:305–311.
36.
Chelius MK, Triplett EW. 2001. The diversity of archaea and bacteria in association with the roots of Zea mays L. Microb Ecol 41:252–263.
37.
Sliwinski MK, Goodman RM. 2004. Comparison of crenarchaeal consortia inhabiting the rhizosphere of diverse terrestrial plants with those in bulk soil in native environments. Appl Environ Microbiol 70:1821–1826.
38.
Wang H, Bier R, Zgleszewski L, Peipoch M, Omondi E, Mukherjee A, Chen F, Zhang C, Kan J. 2020. Distinct distribution of archaea from soil to freshwater to estuary: implications of archaeal composition and function in different environments. Front Microbiol 11:576661.
39.
Liu N, Hu H, Ma W, Deng Y, Liu Y, Hao B, Zhang X, Dimitrov D, Feng X, Wang Z, Lloyd Karen G. 2019. Contrasting biogeographic patterns of bacterial and archaeal diversity in the top- and subsoils of temperate grasslands. mSystems 4:e00566-19.
40.
Leininger S, Urich T, Schloter M, Schwark L, Qi J, Nicol GW, Prosser JI, Schuster SC, Schleper C. 2006. Archaea predominate among ammonia-oxidizing prokaryotes in soils. Nat New Biol 442:806–809.
41.
Wang J-T, Cao P, Hu H-W, Li J, Han L-L, Zhang L-M, Zheng Y-M, He J-Z. 2015. Altitudinal distribution patterns of soil bacterial and archaeal communities along Mt. Shegyla on the Tibetan Plateau. Microb Ecol 69:135–145.
42.
Mamet SD, Lamb EG, Piper CL, Winsley T, Siciliano SD. 2017. Archaea and bacteria mediate the effects of native species root loss on fungi during plant invasion. ISME J 11:1261–1275.
43.
Zhang L-M, Offre PR, He J-Z, Verhamme DT, Nicol GW, Prosser JI. 2010. Autotrophic ammonia oxidation by soil thaumarchaea. Proc Natl Acad Sci U S A 107:17240–17245.
44.
Hatzenpichler R. 2012. Diversity, physiology, and niche differentiation of ammonia-oxidizing archaea. Appl Environ Microbiol 78:7501–7510.
45.
Neill SJ, Desikan R, Hancock JT. 2003. Nitric oxide signalling in plants. New Phytol 159:11–35.
46.
Coskun D, Britto DT, Shi W, Kronzucker HJ. 2017. Nitrogen transformations in modern agriculture and the role of biological nitrification inhibition. Nat Plants 3:17074.
47.
Kalyaanamoorthy S, Minh BQ, Wong TKF, von Haeseler A, Jermiin LS. 2017. ModelFinder: fast model selection for accurate phylogenetic estimates. Nat Methods 14:587–589.
48.
Nguyen L-T, Schmidt HA, von Haeseler A, Minh BQ. 2015. IQ-TREE: a fast and effective stochastic algorithm for estimating maximum-likelihood phylogenies. Mol Biol Evol 32:268–274.
49.
Alves RJE, Minh BQ, Urich T, von Haeseler A, Schleper C. 2018. Unifying the global phylogeny and environmental distribution of ammonia-oxidising archaea based on amoA genes. Nat Commun 9:1517.
50.
Jung M-Y, Kim J-G, Sinninghe Damsté JS, Rijpstra WIC, Madsen EL, Kim S-J, Hong H, Si O-J, Kerou M, Schleper C, Rhee S-K. 2016. A hydrophobic ammonia-oxidizing archaeon of the Nitrosocosmicus clade isolated from coal tar-contaminated sediment. Environ Microbiol Rep 8:983–992.
51.
Liu L, Liu M, Jiang Y, Lin W, Luo J. 2021. Production and excretion of polyamines to tolerate high ammonia, a case study on soil ammonia-oxidizing archaeon “Candidatus Nitrosocosmicus agrestis” mSystems 6:e01003-20.
52.
Sauder LA, Albertsen M, Engel K, Schwarz J, Nielsen PH, Wagner M, Neufeld JD. 2017. Cultivation and characterization of Candidatus Nitrosocosmicus exaquare, an ammonia-oxidizing archaeon from a municipal wastewater treatment system. ISME J 11:1142–1157.
53.
Abby SS, Kerou M, Schleper C. 2020. Ancestral reconstructions decipher major adaptations of ammonia-oxidizing archaea upon radiation into moderate terrestrial and marine environments. MBio 11:e02371-20.
54.
Lehtovirta-Morley LE, Ross J, Hink L, Weber EB, Gubry-Rangin C, Thion C, Prosser JI, Nicol GW. 2016. Isolation of “Candidatus Nitrosocosmicus franklandus”, a novel ureolytic soil archaeal ammonia oxidiser with tolerance to high ammonia concentration. FEMS Microbiol Ecol 92:fiw057.
55.
Wood NJ, Sørensen J. 2001. Catalase and superoxide dismutase activity in ammonia-oxidising bacteria. FEMS Microbiol Ecol 38:53–58.
56.
Yeoh YK, Paungfoo-Lonhienne C, Dennis PG, Robinson N, Ragan MA, Schmidt S, Hugenholtz P. 2016. The core root microbiome of sugarcanes cultivated under varying nitrogen fertilizer application. Environ Microbiol 18:1338–1351.
57.
Simon HM, Dodsworth JA, Goodman RM. 2000. Crenarchaeota colonize terrestrial plant roots. Environ Microbiol 2:495–505.
58.
Ai C, Liang G, Sun J, Wang X, He P, Zhou W. 2013. Different roles of rhizosphere effect and long-term fertilization in the activity and community structure of ammonia oxidizers in a calcareous fluvo-aquic soil. Soil Biol Biochem 57:30–42.
59.
Thion CE, Poirel JD, Cornulier T, De Vries FT, Bardgett RD, Prosser JI. 2016. Plant nitrogen-use strategy as a driver of rhizosphere archaeal and bacterial ammonia oxidiser abundance. FEMS Microbiol Ecol 92:fiw091.
60.
Wattenburger CJ, Gutknecht J, Zhang Q, Brutnell T, Hofmockel K, Halverson L. 2020. The rhizosphere and cropping system, but not arbuscular mycorrhizae, affect ammonia oxidizing archaea and bacteria abundances in two agricultural soils. Agric, Ecosyst Environ, Appl Soil Ecol 151:103540.
61.
Norton J, Ouyang Y. 2019. Controls and adaptive management of nitrification in agricultural soils. Front Microbiol 10:1931.
62.
Zhalnina KV, Dias R, Leonard MT, Dorr de Quadros P, Camargo FAO, Drew JC, Farmerie WG, Daroub SH, Triplett EW. 2014. Genome sequence of Candidatus Nitrososphaera evergladensis from group I.1b enriched from everglades soil reveals novel genomic features of the ammonia-oxidizing archaea. PLoS ONE 9:e101648.
63.
Whittaker JW. 2012. Non-heme manganese catalase--the “other” catalase. Arch Biochem Biophys 525:111–120.
64.
Bayer B, Pelikan C, Bittner MJ, Reinthaler T, Könneke M, Herndl GJ, Offre P, Orsi W. 2019. Proteomic response of three marine ammonia-oxidizing archaea to hydrogen peroxide and their metabolic interactions with a heterotrophic alphaproteobacterium. mSystems 4:e00181-19.
65.
Kim J-G, Park S-J, Sinninghe Damsté JS, Schouten S, Rijpstra WIC, Jung M-Y, Kim S-J, Gwak J-H, Hong H, Si O-J, Lee S, Madsen EL, Rhee S-K. 2016. Hydrogen peroxide detoxification is a key mechanism for growth of ammonia-oxidizing archaea. Proc Natl Acad Sci U S A 113:7888–7893.
66.
Simon HM, Jahn CE, Bergerud LT, Sliwinski MK, Weimer PJ, Willis DK, Goodman RM. 2005. Cultivation of mesophilic soil crenarchaeotes in enrichment cultures from plant roots. Appl Environ Microbiol 71:4751–4760.
67.
Veal EA, Day AM, Morgan BA. 2007. Hydrogen peroxide sensing and signaling. Mol Cell 26:1–14.
68.
Zamocky M, Furtmüller PG, Obinger C. 2008. Evolution of catalases from bacteria to humans. Antioxid Redox Signal 10:1527–1548.
69.
Stone JR, Yang S. 2006. Hydrogen peroxide: a signaling messenger. Antioxid Redox Signal 8:243–270.
70.
Imlay JA. 2008. Cellular defenses against superoxide and hydrogen peroxide. Annu Rev Biochem 77:755–776.
71.
Passardi F, Penel C, Dunand C. 2004. Performing the paradoxical: how plant peroxidases modify the cell wall. Trends Plant Sci 9:534–540.
72.
Tobimatsu Y, Schuetz M. 2019. Lignin polymerization: how do plants manage the chemistry so well? Curr Opin Biotechnol 56:75–81.
73.
Dunand C, Crèvecoeur M, Penel C. 2007. Distribution of superoxide and hydrogen peroxide in Arabidopsis root and their influence on root development: possible interaction with peroxidases. New Phyto 174:332–341.
74.
Shin R, Schachtman DP. 2004. Hydrogen peroxide mediates plant root cell response to nutrient deprivation. Proc Natl Acad Sci U S A 101:8827–8832.
75.
Lee SH, Singh AP, Chung GC. 2004. Rapid accumulation of hydrogen peroxide in cucumber roots due to exposure to low temperature appears to mediate decreases in water transport. J Exp Bot 55:1733–1741.
76.
Zhao Y, Zhang Y, Liu F, Wang R, Huang L, Shen W. 2019. Hydrogen peroxide is involved in methane-induced tomato lateral root formation. Plant Cell Rep 38:377–389.
77.
Liszkay A, van der Zalm E, Schopfer P. 2004. Production of reactive oxygen intermediates (O2 ˙−, H2O2, and ˙OH) by maize roots and their role in wall loosening and elongation growth. Plant Physiol 136:3114–3123.
78.
Huang Y-M, Zou Y-N, Wu Q-S. 2017. Alleviation of drought stress by mycorrhizas is related to increased root H2O2 efflux in trifoliate orange. Sci Rep 7:42335.
79.
Dai H, Wu B, Chen B, Ma B, Chu C. 2022. Diel fluctuation of extracellular reactive oxygen species production in the rhizosphere of rice. Environ Sci Technol 56:9075–9082.
80.
Ikuta S, Matuura K, Imamura S, Misaki H, Horiuti Y. 1977. Oxidative pathway of choline to betaine in the soluble fraction prepared from Arthrobacter globiformis. J Biochem 82:157–163.
81.
Kontchou CY, Blondeau R. 1990. Isolation and characterization of hydrogen peroxide producing Aerococcus sp. from soil samples. FEMS Microbiol Lett 68:323–327.
82.
Herman DJ, Johnson KK, Jaeger CH, Schwartz E, Firestone MK. 2006. Root influence on nitrogen mineralization and nitrification in Avena barbata rhizosphere soil. Soil Sci Soc Am J 70:1504–1511.
83.
Ofek-Lalzar M, Sela N, Goldman-Voronov M, Green SJ, Hadar Y, Minz D. 2014. Niche and host-associated functional signatures of the root surface microbiome. Nat Commun 5:4950.
84.
Jung M-Y, Park S-J, Kim S-J, Kim J-G, Sinninghe Damsté JS, Jeon CO, Rhee S-K. 2014. A mesophilic, autotrophic, ammonia-oxidizing archaeon of thaumarchaeal group I.1a cultivated from a deep oligotrophic soil horizon. Appl Environ Microbiol 80:3645–3655.
85.
Jung M-Y, Park S-J, Min D, Kim J-S, Rijpstra WIC, Sinninghe Damsté JS, Kim G-J, Madsen EL, Rhee S-K. 2011. Enrichment and characterization of an autotrophic ammonia-oxidizing archaeon of mesophilic crenarchaeal group I.1a from an agricultural soil. Appl Environ Microbiol 77:8635–8647.
86.
Jung M-Y, Gwak J-H, Rohe L, Giesemann A, Kim J-G, Well R, Madsen EL, Herbold CW, Wagner M, Rhee S-K. 2019. Indications for enzymatic denitrification to N2O at low pH in an ammonia-oxidizing archaeon. ISME J 13:2633–2638.
87.
Koranda M, Schnecker J, Kaiser C, Fuchslueger L, Kitzler B, Stange CF, Sessitsch A, Zechmeister-Boltenstern S, Richter A. 2011. Microbial processes and community composition in the rhizosphere of European beech - the influence of plant C exudates. Soil Biol Biochem 43:551–558.
88.
Jung M-Y, Sedlacek CJ, Kits KD, Mueller AJ, Rhee S-K, Hink L, Nicol GW, Bayer B, Lehtovirta-Morley L, Wright C, de la Torre JR, Herbold CW, Pjevac P, Daims H, Wagner M. 2022. Ammonia-oxidizing archaea possess a wide range of cellular ammonia affinities. ISME J 16:272–283.
89.
Beeckman F, Motte H, Beeckman T. 2018. Nitrification in agricultural soils: impact, actors and mitigation. Curr Opin Biotechnol 50:166–173.
90.
Neumann G, Römheld V. 2012. Rhizosphere chemistry in relation to plant nutrition, p 347–368. In Marschner P (ed), Marschner’s mineral nutrition of higher plants, 3rd ed. Academic Press.
91.
Xu G, Fan X, Miller AJ. 2012. Plant nitrogen assimilation and use efficiency. Annu Rev Plant Biol 63:153–182.
92.
Song GC, Im H, Jung J, Lee S, Jung M-Y, Rhee S-K, Ryu C-M. 2019. Plant growth-promoting archaea trigger induced systemic resistance in Arabidopsis thaliana against Pectobacterium carotovorum and Pseudomonas syringae. Environ Microbiol 21:940–948.
93.
Awala SI, Gwak J-H, Kim Y-M, Kim S-J, Strazzulli A, Dunfield PF, Yoon H, Kim G-J, Rhee S-K. 2021. Verrucomicrobial methanotrophs grow on diverse C3 compounds and use a homolog of particulate methane monooxygenase to oxidize acetone. ISME J 15:3636–3647.
94.
Nolan T, Hands RE, Ogunkolade W, Bustin SA. 2006. SPUD: a quantitative PCR assay for the detection of inhibitors in nucleic acid preparations. Anal Biochem 351:308–310.
95.
Martin M. 2011. Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnet J 17:10.
96.
Callahan BJ, McMurdie PJ, Rosen MJ, Han AW, Johnson AJA, Holmes SP. 2016. DADA2: high-resolution sample inference from Illumina amplicon data. Nat Methods 13:581–583.
97.
Katoh K, Standley DM. 2013. MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol Biol Evol 30:772–780.
98.
Gwak J-H, Jung M-Y, Hong H, Kim J-G, Quan Z-X, Reinfelder JR, Spasov E, Neufeld JD, Wagner M, Rhee S-K. 2020. Archaeal nitrification is constrained by copper complexation with organic matter in municipal wastewater treatment plants. ISME J 14:335–346.
99.
Shen T, Stieglmeier M, Dai J, Urich T, Schleper C. 2013. Responses of the terrestrial ammonia-oxidizing archaeon Ca. Nitrososphaera viennensis and the ammonia-oxidizing bacterium Nitrosospira multiformis to nitrification inhibitors. FEMS Microbiol Lett 344:121–129.
100.
McMurdie PJ, Holmes S. 2013. Phyloseq: an R package for reproducible interactive analysis and graphics of microbiome census data. PLoS One 8:e61217.
101.
Oksanen J, Blanchet FG, Kindt R, Legendre P, Minchin P, O’hara R, Simpson G, Solymos P, Stevens MHH, Wagner H. 2018. Vegan: community ecology package.
102.
Wickham H. 2016. ggplot2: elegant graphics for data analysis. Springer, New York.
103.
Roberts DW. 2017. labdsv: ordination and multivariate analysis for ecology. R package version 2.0-1
Information & Contributors
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mSphere
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eLocator: e00821-24
Editor: Maria L. Marco, University of California at Davis, Davis, California, USA
PubMed: 39530672
Copyright
© 2024 Lee et al. This is an open-access article distributed under the terms of the Creative Commons Attribution 4.0 International license.
History
Received: 1 October 2024
Accepted: 11 October 2024
Published online: 12 November 2024
Keywords
Data Availability
The 16S rRNA, AOA amoA, and AOA MnKat gene amplicon sequencing data generated in this study have been deposited in NCBI under the BioProject ID PRJNA905906.
Contributors
Editor
Maria L. Marco
Editor
University of California at Davis, Davis, California, USA
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“Ca. Nitrosocosmicus” members are the dominant archaea associated with plant rhizospheres. mSphere
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