Skip to main content
Download PDF

Rhizobiome diversity of field-collected hyperaccumulating Noccaea sp.

BMC Plant Biology volume 24, Article number: 922 (2024) Cite this article

Abstract

Hyperaccumulating plants are able to (hyper)accumulate high concentrations of metal(loid)s in their above-ground tissues without any signs of toxicity. Studies on the root-associated microbiome have been previously conducted in relation to hyperaccumulators, yet much remains unknown about the interactions between hyperaccumulating hosts and their microbiomes, as well as the dynamics within these microbial communities. Here, we assess the impact of the plant host on shaping microbial communities of three naturally occurring populations of Noccaea species in Slovenia: Noccaea praecox and co-occurring N. caerulescens from the non-metalliferous site and N. praecox from the metalliferous site. We investigated the effect of metal enrichment on microbial communities and explored the interactions within microbial groups and their environment. The abundance of bacterial phyla was more homogeneous than fungal classes across all three Noccaea populations and across the three root-associated compartments (roots, rhizosphere, and bulk soil). While most fungal and bacterial Operational Taxonomic Units (OTUs) were found at both sites, the metalliferous site comprised more unique OTUs in the root and rhizosphere compartments than the non-metalliferous site. In contrast to fungi, bacteria exhibited differentially significant abundance between the metalliferous and non-metalliferous sites as well as statistically significant correlations with most of the soil parameters. Results revealed N. caerulescens had the highest number of negative correlations between the bacterial phyla, whereas the population from the metalliferous site had the fewest. This decrease was accompanied by a big perturbation in the bacterial community at the metalliferous site, indicating increased selection between the bacterial taxa and the formation of potentially less stable rhizobiomes. These findings provide fundamentals for future research on the dynamics between hyperaccumulators and their associated microbiome.

Peer Review reports

Background

High concentration of metal(loid)s in the substrate is a ubiquitous abiotic stressor that negatively affects plant growth and development [1]. Plants have evolved different mechanisms to tolerate the excess metal(loid) concentration in the soil, with hyperaccumulating species developing an extraordinary set of traits that enable them to (hyper)accumulate and tolerate high concentrations of one or more metal(loid)s in their aboveground biomass, without any visible signs of toxicity [2, 3]. To date, more than 700 plant species are recognized as hyperaccumulators [4], with several belonging to Brassicaceae [5]. Within this family, a group of hyperaccumulators has been identified in the genus Noccaea, including a model hyperaccumulating species Noccaea caerulescens [6], and a closely related N. praecox [7]. Both species are zinc (Zn) and cadmium (Cd) hyperaccumulators [8,9,10], which is a rare phenomenon; namely, 75% of hyperaccumulators (hyper)accumulate nickel (Ni) [11]. While Zn and Ni are essential elements for plants, Cd and Pb have no known function and are toxic at very low concentrations to the majority of plants [12,13,14].

The structure and the diversity of microbial communities are influenced by several factors, including the type of soil, soil compartments [15,16,17], and plant host [15]. The presence of metals in soil can negatively impact microbial community structure and reduce microbial abundance [18]. The effect of plant host on the root-associated microbiome is especially visible in hyperaccumulating plants, as they can select for metal-tolerant bacteria in the proximity of their roots, even if the plant grows in soil that is not enriched with metals [17]. By contrast, fungi seem to be more influenced by the soil [19, 20] and are more dominantly associated with non-hyperaccumulators than hyperaccumulators growing in the same location [17]. Nevertheless, a combination of plant host and environment significantly shapes the microbial communities [15, 21]. Although hyperaccumulators have a conserved core rhizosphere microbiota [22], the presence of metal(loid)s significantly impacts their microbial community structure in the soil [23, 24]. When growing in metalliferous soils, hyperaccumulators have been shown to harbour a higher number of bacterial and fungal taxa compared to non-metalliferous soils, including specific microbes that help the plant alleviate metal stress and facilitate its growth and survival [21, 25]. Plants actively recruit their microbiome [26]. In particular, hyperaccumulators growing on metalliferous soils have been shown to consistently attract soil microbes that may contribute to phytoextraction [15, 27, 28]. However, the root-associated microbiome of hyperaccumulating plants has been scarcely studied. Therefore, detailed investigations are required in different hyperaccumulating plants to pave the way for a better understanding of the hyperaccumulating traits to fill in the gaps of knowledge in rhizobiome diversity in different hyperaccumulators.

Recent advancements in soil microbial ecology, driven by high-throughput sequencing techniques, have highlighted a new challenge: understanding the intricate relationships between microorganisms. Network models compel us to view taxa not in isolation but as interconnected components within microbial communities and their environment [29]. Microorganisms can establish a range of relationships, such as mutualism, synergism, and antagonism, all of which can be considered positive (e.g., mutualism) and negative (e.g., antagonism). These ecological interactions represent critical evolutionary pressures for natural selection during microbial evolution. As such, network analysis offers profound insights into ecosystem organization, beyond its role as a visualization tool. Nevertheless, careful interpretation of results is crucial because significant spatial associations between species (or their absence) do not necessarily represent true ecological interactions [30, 31].

The multi-omics approaches have the potential to significantly expand and complement our understanding of unique physiological traits found in hyperaccumulators and of the ways they interact with the environment. Here, we report on the root-associated microbiomes of the hyperaccumulating Noccaea species in its natural environment. More specifically, we aim to: (i) estimate the impact of the plant host on the formation of the microbial communities unique to N. praecox and compare it to the microbiome of another hyperaccumulating species, N. caerulescens, growing in the same non-metalliferous location, (ii) identify changes in the microbial communities of N. praecox under metal enrichment compared to N. praecox from the non-metalliferous site, and (iii) estimate the connections within individual microbial groups and with their environment. We hypothesize that the plant host and soil metal status tailor the associated microbial community, thereby forming a specific core community. With this in mind, we: (i) analysed the bacterial and fungal rhizobiomes of the hyperaccumulating N. caerulescens in non-metalliferous soil and of hyperaccumulating N. praecox in metalliferous and non-metalliferous soils, (ii) assessed the adherence to the neutral model to eliminate stochastic processes, and (iii) evaluated the importance of the environment and microbial interactions on the formation of root-associated microbiomes of these two hyperaccumulating species.

Results

Microbial communities associated with Noccaea hyperaccumulators

Microbiome sequencing of N. caerulescens and N. praecox roots with accompanying rhizosphere and bulk soil revealed 2996 and 39 unique OTUs for bacteria and fungi, respectively. PerMANOVA confirmed that soil compartments (i.e., roots, rhizosphere, and bulk soil) harbor statistically different bacterial and fungal communities (Table S2). The α-diversity within the bacterial community showed nearly identical values for rhizosphere and bulk soil. The number of observed OTUs was statistically lower in roots than in rhizosphere and bulk soil, whereas Chao1 richness was statistically higher. In contrast, Shannon index did not show a statistically significant difference between the soil compartments (Table S3). In the fungal community, the number of observed OTUs was the highest in roots and decreased from the root to the rhizosphere and bulk soil. Chao1 richness and Shannon index showed no statistically significant differences between the soil compartments (Table S3). Overall, the diversity and richness of fungi were significantly lower compared to those of bacteria (Table S3). Actinobacteria and Proteobacteria (44% and 35% average relative abundance, respectively) were dominant across all three Noccaea populations, and root-associated compartments (Fig. 1a). Among the fungal community, Ascomycota was the most dominant group, with Dothiomycetes and Leotiomycetes representing on average over 73% of all fungal sequences (Fig. 1b). Notably, arbuscular mycorrhizal fungi (Glomeromycetes) from Archaeosporales and Diversisporales comprised 9% of all fungal sequences. Additionally, Agaricomycetes displayed a significant level of abundance in the rhizosphere of N. praecox from the metalliferous site.

The majority of bacterial phyla indicated no clear preferences for specific soil compartments (Fig. 1c, Suppl. Mat. Fig.S1), and a large majority of bacterial OTUs were found in all three compartments (Fig. 1e, Suppl. Mat. Fig. S1). For individual species/location combination, 1884–2089 bacterial OTUs (69–77% of all bacterial OTUs) were shared between all three soil compartments (Suppl. Mat. Fig.S1). Bulk soil showed the lowest number of unique bacterial OTUs with 43–61 OTUs (2% of all bacterial OTUs). The number of unique OTUs in roots and rhizosphere was similar for both Noccaea species at Lokovec but lower compared to the rhizosphere at the metalliferous site Žerjav (Suppl. Mat. Fig. S1). Similar to the bacterial community, a large portion of fungal OTUs revealed no preference for a particular soil compartment (Fig. 1f, Suppl. Mat. Fig. S2), although the distribution among individual compartments indicated that fungal OTUs preferred the root-rhizosphere environment over the bulk soil (Fig. 1d, Suppl. Mat. Fig. S2). The majority of unique OTUs were observed for the root compartment with 7–13 OTUs (26–52% of all fungal OTUs).

Fig. 1
figure 1

Top ten abundances of (a) bacterial phyla and (b) fungal classes of root and rhizosphere compartments of Noccaea praecox (Np) and N. caerulescens (Nc) and bulk soil from the non-metalliferous (Lo) and metalliferous (Ze) sites; ternary plots indicate the fraction of (c) bacterial families and (d) fungal classes with the top ten abundances in each soil compartment (for all sites and plant species). The symbol size (black circle) represents the average abundance; the Venn diagram shows the number of common and specific Operational Taxonomic Units (OTUs) in (e) bacterial and (f) fungal communities and the proportion of the total number of sequences in each compartment (for all sites and plant species)

Full size image

Effect of metal-enrichment on microbial communities of N. praecox

Next, we compared the microbial communities in N. praecox only, i.e., from metalliferous and non-metalliferous sites. The comparison showed that not only the compartment but also the location is a significant factor that shapes the root-associated microbiome in Noccaea hyperaccumulatos (Table S2 The α-diversity of the bacterial community revealed that the non-metalliferous site in Lokovec has higher diversity and species richness compared to the metalliferous site in Žerjav (Table S4), although differences were not statistically significant. In contrast, the fungal community was more diverse with more species richness in the metalliferous site (Table S4). Again, the sites did not differ significantly. In root and rhizosphere compartments of N. praecox, the majority of fungal and bacterial OTUs identified were found at both sampling sites (Fig. 2). At the metalliferous site in Žerjav, eight unique fungal OTUs (24% of all fungal OTUs) and 262 unique bacterial OTUs (9% of all bacterial OTUs) were found (Fig. 2a), while at the non-metalliferous site in Lokovec six unique fungal OTUs (18% of all OTUs) and 211 unique bacterial OTUs (7% of all OTUs) were found (Fig. 2b).

Fig. 2
figure 2

Venn diagrams of common and unique Operational Taxonomic Units of rhizosphere and root compartments for (a) fungal and (b) bacterial communities of Noccaea praecox at the non-metalliferous (Lokovec) and metalliferous (Žerjav) sites. Numbers represent percentages of all taxa with the absolute numbers in parentheses

Full size image

The follow-up analysis of keystone bacterial OTUs in the root and rhizosphere compartments revealed no shared keystone bacterial OTUs between the two populations of N. praecox (Table 1). In the non-metalliferous site in Lokovec, one keystone OTU (Unclassified Devosiaceae) belonging to the class Alphaproteobacteria was detected, whereas at the metalliferous site in Žerjav, Fimbriiglobus and Phycicoccus, belonging to the classes Planctomycetia and Actinomycetes, respectively, were identified. Notably, the fungal community exhibited a higher number of keystone OTUs compared to bacteria, as outlined in Table 1. Although no common keystone OTUs were detected, Acephala and Golovinomycetes from the control site belong to the class Leotiomycetes, which was found as a keystone OTU in the metalliferous site. Additionally, Thelephora (Agaricomycetes) was found as a keystone OTU in Lokovec, whereas in Žerjav, a keystone belonging to the Dothideomycetes was detected (Unclassified Pleosporales).

Table 1 Presence (indicated by “x”) of keystone bacterial operational taxonomic units (OTUs) in the rhizosphere and root compartments of Noccaea praecox from the non-metalliferous site (Lokovec) and metalliferous site (Žerjav)
Full size table

Measurement of Zn, Cd and Pb root concentrations with ICP-MS showed that root Zn and Cd concentrations were higher in N. praecox than in N. caerulescens regardless of the location, with concentration at metalliferous site reaching 816 ± 87.5 mg kg− 1 and 699 ± 158 mg kg− 1, respectively (Table 2). Root Pb concentrations in N. praecox from Žerjav reached 5161 ± 1242 mg kg− 1, which is over 500 times the concentration in the roots of Noccaea plants at the non-metalliferous site. In contrast, no difference in root Pb concentrations was observed in Noccaea species at the non-metalliferous site in Lokovec.

Table 2 Concentration of zinc (zn), cadmium (cd), and lead (pb) in roots of Noccaea caerulescens and N. praecox naturally occurring on non-metalliferous (Lokovec) or metalliferous (Žerjav) sites. Shown are means ± standard errors (n = 5). Different letters in bold depict statistically significant differences between populations (Dunn’s test at p < 0.05)
Full size table

To evaluate the impacts of the two locations and the plant host on the microbial community, we employed DESeq analysis alongside the Sloan neutral model evaluation. No fungal genera that deviated from the neutral model exhibited differentially significant abundance between both sampling sites. In contrast, according to the Sloan neutral model, the bacterial community included 394 bacterial genera that were above the prediction and 282 genera that were below the prediction. Among these, 92 and 37 bacterial genera were enriched and under-enriched, respectively, and statistically significant according to the DESeq analysis. The highest number of genera not following the neutral model and displaying statistically significant enrichment was observed for Actinobacteria (6 taxa), Bacterioidetes (6 taxa), and Proteobacteria (6 taxa). Proteobacteria also exhibited the highest number of genera that were below prediction (23 taxa), followed by Verrucomicrobia (7 taxa) and Cyanobacteria (6 taxa).

Further, it was explored how fungal and bacterial communities that do not follow the Sloan neutral model correlated with soil properties. Therefore, we employed the Mantel test for rhizosphere and root compartments. Surprisingly, the fungal community exhibited no statistically significant correlation with any of the soil parameters in either compartment. In contrast, the majority of soil parameters were statistically significantly correlated with the bacterial community in both compartments (Fig. 3). Among all soil parameters, only base saturation did not reveal a statistically significant correlation with the bacterial community in either compartment. In addition, root metal concentrations also showed a statistically significant Mantel correlation with the bacterial communities in the roots and rhizosphere compartments.

Fig. 3
figure 3

Mantel test for (a) root and (b) rhizosphere compartments of bacterial community that did not follow the Sloan neutral model evaluation. Color gradient in squares represents Spearman’s correlation coefficient, line color statistical significance, and line width Mantel’s correlation coefficient (n = 9)

Full size image

Bacterial networks of Noccaea rhizobiome

The co-occurrence network analysis revealed that the majority of bacterial phyla in all three compartments form positive interactions within the bacterial community (Fig. 4a, b), with the exception of Firmicutes, Gemmatimonades, and Bacteroidetes.

Fig. 4
figure 4

Co-occurrence networks for (a) top 15 bacterial phyla of all three compartments (bulk soil, rhizosphere, and roots) and all three Noccaea populations with rho > 0.8 and (b) Spearman correlation plots for bacterial phyla in the rhizosphere and root compartments from the individual Noccaea population with (only correlations with p < 0.05 are shown, n = 6). Lokovec is a non-metalliferous and Žerjav is metalliferous site

Full size image

Bacteroides and Firmicutes exhibited several negative interactions with other bacterial phyla in the roots and rhizosphere of both plant species and at both locations. N. praecox population from Lokovec has a higher number of statistically significant correlations within the root and rhizosphere compartments compared to the N. praecox from the metalliferous site. The number of negative interactions between the bacterial phyla was the highest for N. caerulescens (Fig. 4b). In contrast, the overall number of negative edges was the highest for N. praecox at the non-metaliferous site (849 negative edges) and the lowest for N. praecox at the metaliferous site (259 negative edges) (Fig. 5). This change was a result of a severe decrease in negative edges of the dominant bacterial phyla Proteobacteria (from 401 down to 165 negative edges) and Actinobacteria (from 295 down to 38 negative edges) and a slight increase in negative edges in Bacteroides (from 9 up to 27 negative edges) and Firmicutes (from 5 up to 19 negative edges).

The Noccaea population from metalliferous soil had the highest number of bacterial phyla with only positive correlations: Acidobacteria, Candidatus Rokubacteria, Candidatus Tectomicrobia, and Planctomycetes. The other two populations from Lokovec had one phylum each that did not form negative correlations – Acidobacteria and Chloroflexi in N. praecox and N. caerulescens, respectively.

Fig. 5
figure 5

Number of negative edges in bacterial co-occurrence networks for root and rhizosphere compartments of Noccaea populations from the non-metalliferous (Lokovec) and metalliferous (Žerjav) sites. Only correlations with p < 0.05 are presented on the pie charts. Percentages represent the proportion of negative correlations out of the total number of correlations for each phylum, whereas numbers in parentheses present the cumulative sum of the number of negative correlations for each phylum

Full size image

Discussion

Microbial communities associated with Noccaea hyperaccumulators

In our study, root-associated compartments of the hyperaccumulating Noccaea species – bulk soil, rhizosphere, and roots – showed statistically significant differences in abundance and diversity of bacteria and fungi. The α-diversity of the bacterial and fungal communities differed between compartments, indicating that they represent important niches for different microorganisms. In the bacterial community, the diversity was similar between bulk soil and rhizosphere, with a significant decrease in the root compartment, whereas fungal community diversity declined from the root compartment to bulk soil. Other authors report similar observations for bacterial [15] and fungal communities [32]. Species richness, determined by the Chao1 index, was the highest for the root compartment for both bacterial and fungal communities, indicating that the host’s roots harbor a higher number of different genera compared to the other two (soil) compartments studied. While fungi indicated a preference for the root-rhizosphere compartments, bacteria did not exhibit a clear preference for any particular soil compartment. This aligns with the results observed by Xiao et al. (2020) [32] in their study on hyperaccumulating fern Pteris vittata. Nevertheless, in roots, Glomeromycetes were significantly more abundant in N. praecox from the metalliferous site in Žerjav compared to both Noccaea populations from the non-metalliferous site in Lokovec. Glomeromycetes are known to form beneficial associations with plants through arbuscular mycorrhiza [33], playing an important role in the diversification and stability of soil microbiota [34]. They are also found in environments rich in metals [35, 36], explaining their presence in Žerjav. However, there is a common belief that Brassicaceae, which includes the genus Noccaea, do not form mycorrhizal associations [37]. Despite this, several studies using both morphological observations as well as molecular have confirmed their presence in the roots of representatives from this family, including N. praecox [8, 38,39,40].

Actinobacteria and Proteobacteria dominance over other bacterial phyla, independent of the soil metal status seems to be a common pattern. Both phyla were also prevalent in other hyperaccumulating species [15, 17] and are positively correlated with Cd/Zn hyperaccumulation in plants [28]. Actinobacteria are known to promote metal absorption and plant growth [41, 42] and are well-represented soil microorganisms [43, 44]. They were also shown to contribute to the increased plant tolerance to heavy metals [45]. Proteobacteria, on the other hand, are also known for their plant growth-promoting effects and resistance to some metals, commonly found on both metalliferous and non-metalliferous sites [46, 47]. Thus, our findings suggest that the studied Noccaea hyperaccumulators are associated with potentially beneficial organisms that could positively influence plant growth and fitness. Interestingly, no differences in the abundances of bacterial genera were observed between the populations from Lokovec, suggesting that the plant host plays a less important role in shaping the microbiota than the location itself.

Effect of metal-enrichment on microbial communities of N. praecox

In addition to compartments, soil metal enrichment was found to be an important factor in shaping the root-associated microbiome. The concentration of Zn, Cd, and Pb were the highest in the roots of N. praecox growing at the metalliferous site, forming an additional selective pressure on the microbial community. In line with this, we observed more unique bacterial OTUs than the non-metalliferous site and a lower α-diversity and species richness in Žerjav compared to Lokovec. Additionally, the uniqueness of fungal genera was slightly higher in Žerjav than in Lokovec, but the difference was not as pronounced as for bacteria, possibly due to the lower number of fungal to bacterial genera detected. Nevertheless, most bacterial and fungal genera were present on both sampling sites. This could be explained by similar soil conditions, i.e., pH, organic matter content, and cation exchange capacity. These soil parameters also influence the bioavailability of metals. A neutral pH combined with a high organic matter content and a high cation exchange capacity increases the adsorption of metals to soil particles and decreases the bioavailability of metals. Conversely to the bacterial community, the α-diversity and species richness of the fungal community were higher in metalliferous compared to non-metalliferous sites. These results partially align with other studies on fungi [21], indicating that polluted sites have significantly higher microbial diversity than non-metalliferous sites [48, 49].

No bacterial keystone taxa of N. praecox were characteristic for the two locations. Phycicoccus and Fimbriiglobus were found as keystone OTUs in the population from Žerjav, whereas unclassified Devosiaceae were found only in N. praecox from Lokovec. Both Phycioccocus and Fimbriiglobus are known to be resistant to environmental contaminants such as heavy metals [50, 51]. Devosiaceae, on the other hand, prefer non-metalliferous environments and are mostly associated with the root compartment [52, 53]. These results suggest the environment is a major factor in determining bacteria that present the core of the local ecosystem. Conversely, the fungal community was more similar between locations, with Leotiomycetes being keystone taxa in both Lokovec and Žerjav, although due to a lack of lower taxonomic level information, it remains unknown whether any keystone species from both locations are the same biological species. Not negligibly, Leotiomycetes were one of the most abundant fungal classes in our study. Pleosporales, which are known for being resistant to metals [54], were found as keystone taxa in metalliferous sites. At Lokovec, Acephala and Golovinomyces (both Leotiomycetes) were detected along with Thelephora basidiomycete. Interestingly, Acephala and Thelephora are fungal genera with well-documented tolerance to metal pollution [55,56,57]. Acephala is classified as a dark septate endophyte (DSE) [58], whereas Thelephora is an ectomycorrhizal fungus [59]. Although ectomycorrhiza is not formed by Noccaea, the presence of these fungi in the rhizosphere could alter the soil conditions in favour of the plant. Indeed, the abundance of Thelephora was even higher in Žerjav than in Lokovec. Nevertheless, we cannot rule out opportunistic commensalism due to root exudate attraction. In addition, a low number of unique fungal genera was observed in our study, hindering straightforward conclusions. Nevertheless, the low diversity of fungal taxa could well be the state of the norm for Brassicaceae, which contain many anti-fungal compounds [60] and are believed to have lost the ability to form typical symbiotic associations with beneficial fungi [33]. A study of endophytes of the related Microthlaspi (Brassicaceae) yielded a wide range of OTUs per population (109–272 OTUs per population) [61], with a lower range just above the numbers observed in our study.

The identified bacterial keystone taxa followed Sloan neutral model, which is in accordance with Wang et al. (2021) [62], who examined microbial communities in a freshwater river continuum in subtropical China. This would suggest that bacterial keystones as module hubs of the communities were mostly neutrally distributed generalists with high abundances and were beneficial to many related OTUs. This was not the case with fungal keystone taxa that mainly deviated from the neutral model. It seems that fungi are more intimately connected to the plant host, resulting in different responses to the environment when compared to bacteria.

To evaluate the stochastics within microbial communities, we applied the Sloan neutral model to assess the distribution of microorganisms associated with N. praecox. The bacterial community revealed the statistical significance of genera that were above and below the prediction according to the Sloan neutral model. This suggests certain bacterial genera are either favored by the plant host and/or environmental conditions or associated with active migration and negative interactions within the community. The fungal community also contained the genera that did not follow the Sloan neutral model, but there was no statistical significance in their abundances between both sampling sites. It indicates that these genera follow the dynamics of passive dispersal [63], possibly by animals, precipitation, or other abiotic and/or biotic vectors. Moreover, host-associated microbial community variation does not come only from plant hosts or associated microorganisms. Neutral processes like drift and dispersal are powerful enough to create a large amount of diversity within and among hosts [64] or environmental conditions [65], explaining a significant part of the microbial community structure (64).

Eliminating the randomness of the bacterial community allowed us to evaluate the influence of individual abiotic soil parameters and biotic interactions on the composition of the bacterial OTUs that did not follow the Sloan neutral model. As fungi were less connected to the environment, only bacterial OTUs showed statistically significant correlations with environmental parameters, with the majority affecting the composition of the bacterial community. The root compartment displayed a pattern similar to that of the rhizosphere, suggesting that the environment impacts microbial taxa that colonize the root endosphere or at least the rhizoplane. However, there is still no consensus on how much environmental conditions affect the fungal communities, as different studies report different results [19, 20, 66, 67] In contrast, it is well established that soil bacteria are much more susceptible to biotic and abiotic factors [68, 69] than fungi, which aligns with our results.

Bacterial networks of Noccaea rhizobiome

Along with abiotic environmental factors, microbial interactions also shape the structure and function of the microbiome [70, 71]. The predominance of positive interactions over negative edges indicates a prevalence of cooperative or syntrophic interactions with the potential for extensive mutualistic interactions [28, 72] and is probably a result of heterogeneous microenvironments that reduce direct competition [73].

However, we observed a few negative interactions associated with Gemmatimonadetes, Firmicutes, and Bacteroidetes, indicating potential antagonistic behaviors, possibly arising from competition for the same ecological niche among these microbial entities. Most negative correlations in all three Noccaea populations were associated with Firmicutes, which are a common bacterial phylum involved in various processes, including the decomposition of organic matter and nutrient cycling, and thus important for the maintenance of the soil ecosystem [74, 75]. Firmicutes are highly resistant to toxic metals, and the high percentage of negative correlations could reflect their high competitiveness, as they can displace other phyla in metalliferous soil [76]. Similarly, Bacteroidetes are another metal-resistant bacterial phylum known for their rapid growth and ability to adapt to various environments, including metal-polluted soils [77,78,79]. They are successful competitors [80] and are known to produce antimicrobial compounds that regulate population-level interactions [81, 82], which might explain negative correlations towards other bacteria within the community. Interestingly, no negative edges were associated with Acidobacteria that were observed to carry a substantial proportion of negative edges in several environments, including soils [73].

We further compared the prevalence of negative edges among bacterial phyla in the root and rhizosphere compartments between both studied locations and Noccaea species. Proteobacteria and Actinobacteria exhibited the highest number of negative correlations. Notably, they were also the most abundant phyla within the bacterial community, allowing them to compete for their root-associated niche with other bacterial phyla efficiently [28]. Nevertheless, the number of negative edges in Proteobacteria and Actinobacteria decreased dramatically at the metalliferous site and increased in Firmicutes and Bacterioidetes at the metalliferous site. The overall presence of negative edges inside the community can be an important factor, as they are hypothesized to promote community stability due to negative feedback [83]. In contrast, positive interactions may destabilize the community through the generation of co-dependencies [83]. N. praecox growing at the metalliferous site exhibited Zn, Cd, and Pb root concentrations that were up to 10-times, 76-times, and 737-times those at the non-metalliferous site, respectively, thus increasing the environmental pressure and selection of bacterial groups. In this context, the results of network analyses suggest that conditions at the metalliferous site may increase competition in some bacterial phyla, as indicated by an increased number of negative edges within Firmicutes and Bacterioidetes. However, the overall frequency of these interactions is significantly reduced due to the negative selection of dominant bacterial groups, which could potentially lead to a more unstable community.

Nevertheless, the network analysis included three samples of each soil compartment for the individual soil/habitat combinations, which could introduce some bias into the results. To strengthen the generalizability of these findings, further research with a larger sample size is recommended.

Conclusions

Our study revealed differences in bacterial and fungal communities from Noccaea plants growing at polluted and unpolluted sites, as well as in root-associated compartments. While fungal OTUs varied among populations and compartments, bacterial OTUs displayed more consistent abundance and significant correlations with soil parameters, particularly in the metalliferous site. N. caerulescens exhibited the highest number of negative correlations between the bacterial phyla, while N. praecox population from the metalliferous site the fewest, suggesting microbial specialization or reduced competition. However, the evaluation showed a big perturbation in the bacterial community at the metalliferous site, suggesting increased selection between the bacterial taxa and the formation of potentially less stable rhizobiomes. These results offer valuable insights for future research on hyperaccumulator-microbiome dynamics.

Methods

Site description

Sampling took place at two sites in Slovenia where Noccaea species occur naturally: in Lokovec (N 46° 2′ 39.2706″, E 13° 46′ 8.9934″), a non-metalliferous site where both species co-occur, and in Žerjav (N 46° 28′ 26.1258″, E 14° 51′ 56.0118″), a metal-polluted site where only N. praecox grows. The soils at both sites belong to the reference group of Phaeozems with calcareous rocks, namely limestone in Lokovec and dolomite in Žerjav, as the parent material. Both soils are eutric, with a pH of 6.8 and 6.9 for Lokovec and Žerjav, respectively, and a high base saturation. Among the exchangeable basic cations, calcium cations predominate 30.2–52.1 mmolc 100 g− 1, followed by magnesium 2.9–9.3 mmolc 100 g− 1 and potassium 0.39–0.61 mmolc 100 g− 1. Exchangeable sodium is negligible in both soils. Both soils have a high cation exchange capacity, which is due to the high organic matter content, which is 17.1 and 20.2% for Lokovec and Žerjav, respectively (Table S1). The soils differ in soil texture and carbonates content. The high proportion of sand and carbonates in Žerjav is the result of the typical physical weathering of dolomite. On the other hand, the chemical weathering of limestone leads to a more clayey texture and the dissolution of carbonates (Table S1).

Sample collection

Three replicates of flowering plants, including their rhizosphere and the surrounding soil, were dug out in spring 2022: both species in Lokovec and N. praecox in Žerjav. The plant material was formally identified by Matevž Likar. In addition, at each sampling site, we took a composite soil sample prepared from ten subsamples of the topsoil (0–10 cm). Subsamples were collected across the entire sampling area, tightly packed in a plastic bag to prevent potential cross-contamination, and taken to the laboratory. Before combining and thoroughly mixing the soil subsamples from each location into a single composite sample, we removed the top organic layer containing dry plant material.

During the transfer, plants were kept in plastic pots and were processed the next day, when the roots, the rhizosphere soil, and the bulk soil were separated. Rhizosphere soil was collected by gently shaking the plant to gather the soil adhering to the roots [84], while the roots were thoroughly washed with tap water, followed by distilled water, and finally dried with paper towels. Water and equipment were sterilized by autoclaving for 30 min at 121 °C. All procedures followed appropriate sterile conditions required to prevent contamination of the material. Subsequently, all samples were stored at -80 °C until further use. With this approach we separated the collected microbial communities into three compartments: (i) roots that include endophytes, epiphytes, and the microbes inhabiting the nearest soil particles; (ii) rhizosphere that includes only microbes not in contact with plant roots and therefore less influenced by the root exudates; (iii) bulk soil that was well removed from the plant roots.

Soil analysis

The soil samples were air-dried, crushed, and sieved through a 2 mm sieve (SIST ISO 11464, 2006). For the metal analyses, the soil samples were additionally crushed and sieved through a 160 μm sieve. The soil analysis procedures were carried out as follows: soil texture was determined using a sedimentation pipette method (SIST ISO 11277, 2011). For soil pH measurement, a 1:5 (v/v) ratio of soil and 0.01 M CaCl2 suspension was employed (SIST ISO 10390, 2006). Total carbon (Ctotal) and nitrogen (N) were determined through dry combustion (ISO 10694, 1995) using an elemental analyzer (Elementar Vario MAX Instrument, Germany). Carbonates content (Carbonates) was determined volumetrically using Scheibler calcimeter following soil reaction with hydrochloric acid (SIST ISO 10693, 1995). The inorganic carbon was calculated from the content of carbonates to further assess organic carbon (Corg), which is approximated as the difference between the total carbon (Ctotal) and the inorganic carbon [85]. Soil organic matter content (Organic matter) was calculated as the product of organic carbon × 1.724 [85], and the ratio between organic carbon and total nitrogen was also determined (Corg/N). Plant-available phosphorus (P available) and potassium (K available) were measured post-extraction with ammonium lactate solution (ÖNORM L 1087, 1993) – phosphorus spectrophotometrically with the development of a blue color and potassium using the FAAS method. The cation exchange capacity (CEC) was determined by a two-step extraction procedure: first by extraction with ammonium acetate (pH = 7), whereby the concentrations of the basic cations (Ca2+, Mg2+, K+, and Na+) were measured, and then by extraction with potassium chloride (KCl). The sum of base cations (SUM cations) was measured by FAAS after the first extraction. Base saturation (Base saturation) was then calculated as the sum of base cations (Ca2+, Mg2+, K+, and Na+) divided by the total CEC. Pseudototal concentrations of Pb, Cd, and Zn were determined by FAAS after digestion with aqua regia in a microwave oven system (SIST ISO 11047:1999).

Each soil parameter was measured three times. Mean values and standard deviations were calculated and are presented in Table S1.

Inductively coupled plasma mass spectrometry (ICP-MS)

The concentrations of Zn, Cd, and Pb in dried roots (n = 5–6) were determined following acid digestion by inductively coupled plasma mass spectrometry (ICP-MS). Dried samples were milled to a fine powder using a mortar and a pestle and accurately weighed powdered material was digested in closed vessels using a microwave digester (MARS Xpress, CEM Microwave Technology, Italy). Samples were digested with 6 mL of concentrated HNO3 and 2 mL of 30% H2O2 at 180 °C (holding for 60 min). Digested samples were diluted to 30 mL with MilliQ water before element analyses. Total Zn, Cd, and Pb concentrations of digested samples were determined by ICP-MS (Nexion 1000, PerkinElmer, Waltham, MA, USA). Blank digestions were performed to determine background concentrations of elements, and a tomato leaf standard (Reference 1573a; National Institute of Standards and Technology, NIST, Gaithersburg, MD, USA) was used as an analytical control [86].

Metagenomics

Whole-community DNA from rhizosphere and bulk soil was extracted using the DNeasy® PowerSoil® Pro Kit (Qiagen) and total genomic DNA from roots using the GenElute™ Plant Genomic DNA Miniprep Kit (Sigma Aldrich) following the manufacturer’s instructions. All DNA samples were stored at -80 °C until sequencing. The DNA quality control and shotgun metagenomic sequencing were conducted by Macrogen company using the Illumina HiSeqX platform (2 × 150 pair-ends) in accordance with the manufacturer’s guidelines, employing TruSeq DNA kit (Illumina). Altogether, 24 samples were sequenced: three samples from rhizosphere and plant roots and two bulk soil samples for each population, yielding on average 29,776,694 ± 6,228,348 reads/sample. For the taxonomic assignment of metagenomic reads, SqueezeMeta [86] pipeline that implements a fast-last common ancestor (LCA) algorithm was used to analyze each query gene hit results as the Diamond [87] search query against the GenBank nr database. Operational taxonomic units (OTUs) were defined at the level of genus.

Bioinformatics and statistical analysis

All statistical analyses were performed in R (v4.3.2). Metal concentrations in plant roots and microbial diversity measures were compared using the Kruskal-Wallis test and Dunn’s test for multiple comparisons. Microbial communities were analyzed using the phyloseq library (v1.46.0) [88], while the results were visualized using the ggplot2 (v3.5.0) or additional R libraries specified below. Before any analyses, abundances were normalized to median sequencing depth to eliminate any bias due to differences in the sampling sequencing depth. Differences between the grouping parameters (compartment, location, and Noccaea population) were tested for all bacterial and fungal OTUs with permutational analysis of variance [89] perMANOVA), using adonis2 function in the vegan (v2.6-6.1) library with 1000 permutations. Differentially abundant OTUs were identified using DESeq analysis, DESeq library (v1.44.0). Only OTUs with padj < 0.05 and − 1 < log2FC > 1 were flagged as differentially abundant between comparisons. The Sloan neutral model for fungal and bacterial communities was assessed according to the protocol outlined by Likar et al. (2023) [65]. Calculations of 95% confidence intervals around for the model-calculated frequencies were done by bootstrapping with 1000 bootstrap replicates. OTUs were sorted into three partitions depending on whether they occurred more frequently than (‘above’ partition), less frequently than (‘below’ partition), or within (‘neutral’ partition) the 95% confidence interval of the neutral model predictions. Only microbial genera that did not adhere to the neutral model were used for evaluation of the effects of plant host and abiotic environment on the microbial community.

Libraries ggcor (v0.9.4) and vegan (v2.6-4) were used to conduct the Mantel test for bacterial communities on root and rhizosphere compartments (n = 9). To evaluate the correlation between compartments and soil parameters, Spearman and Mantel correlation coefficients were applied, assessing the statistical significance between parameters. We ensured that the data followed all the assumptions for Spearman’s correlation, e.i. ordinality, paired observations, and monotonic relationship.

The co-occurrence network was inferred based on Spearman’s Rho between the pairwise OTUs matrix constructed by the R psych (v2.1.9) library. The p-values for multiple testing were calculated using the false discovery rate (FDR) controlling procedure. A valid co-occurrence event was considered robust if the correlation coefficient r >|0.8| and if it was statistically significant at p < 0.05. To reduce network complexity, only the top 15 bacterial phyla were included (Fig. 4a). For the triangular correlation plots of the rhizosphere and root compartments of each Noccaea population (n = 6) only data with a p < 0.05 was considered. R libraries chorddiag (v0.1.3) and corrplot (0.92) were used for visualization.

Keystone OTUs were calculated using phylosmith (v1.0.7), igraph (v2.0.3), and microbiome libraries (v1.24.). For the fungal community, all OTUs were included in the keystone analysis, whereas for the bacterial community, only OTUs with more than 500 reads with a statistical significance p < 0.05 were retained. We identified keystone OTUs separately for the generated networks and defined them as those nodes within the top 1% of node degree values for each network [90, 91].

The analysis of negative edges for bacterial phyla was conducted using igraph (v2.0.3), tidygraph (v. 1.3.1), and microbiome (v. 1.24) libraries. Negative edges were analysed for the top 100 most abundant bacterial genera in the rhizosphere and root compartments, and genera with statistical significance of p < 0.05 were included in the analysis.

Data availability

The raw DNA-seq data have been deposited in the National Center for Biotechnology Information (NCBI) under Sequence Read Archive (SRA) study accession SRR28678510-SRR28678533. The datasets analysed during the current study are available in the Zenodo repository, https://doi.org/10.5281/zenodo.10973096.

Abbreviations

OTUs:

Operational Taxonomic Units

Zn:

Zinc

Cd:

Cadmium

Pb:

Lead

References

  1. Nagajyoti PC, Lee KD, Sreekanth TVM. Heavy metals, occurrence and toxicity for plants: a review. Environ Chem Lett. 2010;8(3):199–216.

    Article  CAS  Google Scholar 

  2. Brooks RR, Lee J, Reeves RD, Jaffre T. Detection of nickeliferous rocks by analysis of herbarium specimens of indicator plants. J Geochem Explor. 1977;7:49–57.

    Article  CAS  Google Scholar 

  3. Rascio N. Metal accumulation by some plants growing on zinc-mine deposits. Oikos. 1977;29(2):250–3.

    Article  CAS  Google Scholar 

  4. Reeves RD, Baker AJM, Jaffré T, Erskine PD, Echevarria G, van der Ent A. A global database for plants that hyperaccumulate metal and metalloid trace elements. New Phytol. 2018;218(2):407–11.

    Article  PubMed  Google Scholar 

  5. van der Ent A, Baker AJM, Reeves RD, Pollard AJ, Schat H. Hyperaccumulators of metal and metalloid trace elements: facts and fiction. Plant Soil. 2013;362(1–2):319–34.

    Google Scholar 

  6. Assunção AGL, Schat H, Aarts MGM. Thlaspi caerulescens, an attractive model species to study heavy metal hyperaccumulation in plants. New Phytol. 2003;159(2):351–60.

    Article  PubMed  Google Scholar 

  7. Likar M, Pongrac P, Vogel-Mikuš K, Regvar M. Molecular diversity and metal accumulation of different Thlaspi praecox populations from Slovenia. Plant Soil. 2010;330(1):195–205.

    Article  CAS  Google Scholar 

  8. Vogel-Mikuš K, Drobne D, Regvar M, Zn. Cd and pb accumulation and arbuscular mycorrhizal colonisation of pennycress Thlaspi praecox Wulf. (Brassicaceae) from the vicinity of a lead mine and smelter in Slovenia. Environ Pollut. 2005;133(2):233–42.

    Article  PubMed  Google Scholar 

  9. Reeves RD, Schwartz C, Morel JL, Edmondson J. Distribution and metal-accumulating behavior of Thlaspi caerulescens and associated metallophytes in France. Int J Phytorem. 2001;3(2):145–72.

    Article  CAS  Google Scholar 

  10. Pongrac P, Zhao FJ, Razinger J, Zrimec A, Regvar M. Physiological responses to cd and zn in two Cd/Zn hyperaccumulating Thlaspi species. Environ Exp Bot. 2009;66(3):479–86.

    Article  CAS  Google Scholar 

  11. Rascio N, Navari-Izzo F. Heavy metal hyperaccumulating plants: how and why do they do it? And what makes them so interesting? Plant Sci. 2011;180(2):169–81.

    Article  CAS  PubMed  Google Scholar 

  12. Hamzah Saleem M, Usman K, Rizwan M, Al Jabri H, Alsafran M. Functions and strategies for enhancing zinc availability in plants for sustainable agriculture. Front Plant Sci. 2022;13:1–13.

    Article  Google Scholar 

  13. Silva EB, Alves IS, Alleoni LRF, Grazziotti PH, Farnezi MMM, Santos LL, et al. Availability and toxic level of cadmium, lead and nickel in contaminated soils. Commun Soil Sci Plant Anal. 2020;51(10):1341–56.

    Article  CAS  Google Scholar 

  14. White PJ, Brown PH. Plant nutrition for sustainable development and global health. Ann Bot. 2010;105(7):1073–80.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  15. Luo J, Tao Q, Wu K, Li J, Qian J, Liang Y, et al. Structural and functional variability in root-associated bacterial microbiomes of Cd/Zn hyperaccumulator Sedum Alfredii. Appl Microbiol Biotechnol. 2017;101(21):7961–76.

    Article  CAS  PubMed  Google Scholar 

  16. Bulgarelli D, Rott M, Schlaeppi K, van Ver Loren E, Ahmadinejad N, Assenza F, et al. Revealing structure and assembly cues for Arabidopsis root-inhabiting bacterial microbiota. Nature. 2012;488(7409):91–5.

    Article  CAS  PubMed  Google Scholar 

  17. Martos S, Busoms S, Pérez-Martín L, Llugany M, Cabot C, Poschenrieder C. Identifying the specific root microbiome of the hyperaccumulator Noccaea Brachypetala growing in non-metalliferous soils. Front Microbiol. 2021;12:1–17.

    Article  Google Scholar 

  18. Kamal S, Prasas R, Varma A. Soil microbial diversity in relation to heavy metals. In: Sherameti I, Varma A, editors. Soil heavy metals. Heidelberg: Springer; 2009. pp. 31–63.

    Google Scholar 

  19. Hogan JA, Jusino MA, Smith ME, Corrales A, Song X, Hu Y, et al. Root-associated fungal communities are influenced more by soils than by plant-host root traits in a Chinese tropical forest. New Phytol. 2023;238(5):1849–64.

    Article  CAS  PubMed  Google Scholar 

  20. Bonito G, Reynolds H, Robeson MS, Nelson J, Hodkinson BP, Tuskan G, et al. Plant host and soil origin influence fungal and bacterial assemblages in the roots of woody plants. Mol Ecol. 2014;23(13):3356–70.

    Article  PubMed  Google Scholar 

  21. Kushwaha P, Neilson JW, Maier RM, Babst-Kostecka A. Soil microbial community and abiotic soil properties influence Zn and Cd hyperaccumulation differently in Arabidopsis halleri. Sci Total Environ. 2022;803:1–12.

    Article  Google Scholar 

  22. Luo J, Gu S, Guo X, Liu Y, Tao Q, Zhao HP, et al. Core Microbiota in the rhizosphere of heavy metal accumulators and its contribution to plant performance. Environ Sci Technol. 2022;56(18):12975–87.

    Article  CAS  PubMed  Google Scholar 

  23. Xu Y, Seshadri B, Bolan N, Sarkar B, Ok YS, Zhang W, et al. Microbial functional diversity and carbon use feedback in soils as affected by heavy metals. Environ Int. 2019;125:478–88.

    Article  CAS  PubMed  Google Scholar 

  24. Tipayno SC, Truu J, Samaddar S, Truu M, Preem JK, Oopkaup K, et al. The bacterial community structure and functional profile in the heavy metal contaminated paddy soils, surrounding a nonferrous smelter in South Korea. Ecol Evol. 2018;8(12):6157–68.

    Article  PubMed  PubMed Central  Google Scholar 

  25. Klimek B, Stępniewska K, Seget B, Pandey VC, Babst-Kostecka A. Diversity and activity of soil biota at a post-mining site highly contaminated with zn and cd are enhanced by metallicolous compared to non-metallicolous Arabidopsis halleri ecotypes. Land Degrad Dev. 2023;34(5):1538–48.

    Article  PubMed  Google Scholar 

  26. Abedini D, Jaupitre S, Bouwmeester H, Dong L. Metabolic interactions in beneficial microbe recruitment by plants. Curr Opin Biotechnol. 2021;70:241–7.

    Article  CAS  PubMed  Google Scholar 

  27. Luo J, Guo X, Tao Q, Li J, Liu Y, Du Y, et al. Succession of the composition and co-occurrence networks of rhizosphere microbiota is linked to Cd/Zn hyperaccumulation. Soil Biol Biochem. 2021;153:1–12.

    Article  Google Scholar 

  28. Wu Y, Santos SS, Vestergård M, Martín González AM, Ma L, Feng Y, et al. A field study reveals links between hyperaccumulating Sedum plants-associated bacterial communities and Cd/Zn uptake and translocation. Sci Total Environ. 2022;805:1–10.

    Article  Google Scholar 

  29. Guseva K, Darcy S, Simon E, Alteio LV, Montesinos-Navarro A, Kaiser C. From diversity to complexity: microbial networks in soils. Soil Biol Biochem. 2022;169:1–19.

    Article  Google Scholar 

  30. Barner AK, Coblentz KE, Hacker SD, Menge BA. Fundamental contradictions among observational and experimental estimates of non-trophic species interactions. Ecology. 2018;99(3):557–66.

    Article  PubMed  Google Scholar 

  31. Blanchet FG, Cazelles K, Gravel D. Co-occurrence is not evidence of ecological interactions. Ecol Lett. 2020;23(7):1050–63.

    Article  PubMed  Google Scholar 

  32. Xiao E, Cui J, Sun W, Jiang S, Huang M, Kong D, et al. Root microbiome assembly of As-hyperaccumulator Pteris vittata and its efficacy in arsenic requisition. Environ Microbiol. 2021;23(4):1959–71.

    Article  CAS  PubMed  Google Scholar 

  33. Ferrol N, Tamayo E, Vargas P. The heavy metal paradox in arbuscular mycorrhizas: from mechanisms to biotechnological applications. J Exp Bot. 2016;67(22):6253–565.

    Article  CAS  PubMed  Google Scholar 

  34. van der Heijden MGA, Klironomos JN, Ursic M, Moutoglis P, Streitwolf-Engel R, Boller T, et al. Mycorrhizal fungal diversity determines plant biodiversity ecosystem variability and productivity. Nature. 1998;396:69–72.

    Article  Google Scholar 

  35. Suárez JP, Herrera P, Kalinhoff C, Vivanco-Galván O, Thangaswamy S. Generalist arbuscular mycorrhizal fungi dominated heavy metal polluted soils at two artisanal and small – scale gold mining sites in southeastern Ecuador. BMC Microbiol. 2023;23(42):1–12.

    Google Scholar 

  36. Sánchez-Castro I, Gianinazzi-Pearson V, Cleyet-Marel JC, Baudoin E, van Tuinen D. Glomeromycota communities survive extreme levels of metal toxicity in an orphan mining site. Sci Total Environ. 2017;598:121–8.

    Article  PubMed  Google Scholar 

  37. Glenn MG, Chew FS, Williams PH. Influence of glucosinolate content of Brassica (Cruciferae) roots on growth of vesicular–arbuscular mycorrhizal fungi. New Phytol. 1988;110(2):217–25.

    Article  CAS  Google Scholar 

  38. Trautwig AN, Jackson MR, Kivlin SN, Stinson KA. Reviewing ecological implications of mycorrhizal fungal interactions in the Brassicaceae. Front Plant Sci. 2023;14:1–12.

    Article  Google Scholar 

  39. Pongrac P, Vogel-Mikuš K, Kump P, Nečemer M, Tolrà R, Poschenrieder C, et al. Changes in elemental uptake and arbuscular mycorrhizal colonisation during the life cycle of Thlaspi praecox Wulfen. Chemosphere. 2007;69(10):1602–9.

    Article  CAS  PubMed  Google Scholar 

  40. Pongrac P, Sonjak S, Vogel-Mikuš K, Kump P, Nečemer M, Regvar M. Roots of metal hyperaccumulating population of Thlaspi praecox (Brassicaceae) harbour arbuscular mycorrhizal and other fungi under experimental conditions. Int J Phytorem. 2009;11(4):347–59.

    Article  CAS  Google Scholar 

  41. Lin L, Guo W, Xing Y, Zhang X, Li Z, Hu C, et al. The actinobacterium Microbacterium sp. 16SH accepts pBBR1-based pPROBE vectors, forms biofilms, invades roots, and fixes N2 associated with micropropagated sugarcane plants. Appl Microbiol Biotechnol. 2012;93(3):1185–95.

    Article  CAS  PubMed  Google Scholar 

  42. Visioli G, D’egidio S, Sanangelantoni AM. The bacterial rhizobiome of hyperaccumulators: future perspectives based on omics analysis and advanced microscopy. Front Plant Sci. 2015;5:1–12.

    Article  Google Scholar 

  43. Yang Y, Wang N, Guo X, Zhang Y, Ye B. Comparative analysis of bacterial community structure in the rhizosphere of maize by high-throughput pyrosequencing. PLoS ONE. 2017;12(5):e0178425.

    Article  PubMed  PubMed Central  Google Scholar 

  44. Lauber CL, Hamady M, Knight R, Fierer N. Pyrosequencing-based assessment of soil pH as a predictor of soil bacterial community structure at the continental scale. Appl Environ Microbiol. 2009;75(15):5111–20.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  45. El Baz S, Baz M, Barakate M, Hassani L, El Gharmali A, Imziln B. Resistance to and accumulation of heavy metals by actinobacteria isolated from abandoned mining areas. Sci World J. 2015;2015:1–14.

    Google Scholar 

  46. Zhao X, Huang J, Zhu X, Chai J, Ji X. Ecological effects of heavy metal pollution on soil microbial community structure and diversity on both sides of a river around a mining area. Int J Environ Res Public Health. 2020;17(5680):1–18.

    Google Scholar 

  47. Pereira LB, Vicentini R, Ottoboni LMM. Changes in the bacterial community of soil from a neutral mine drainage channel. PLoS ONE. 2014;9(5):e96605.

    Article  PubMed  PubMed Central  Google Scholar 

  48. Dong Y, Wu S, Deng Y, Wang S, Fan H, Li X, et al. Distinct functions and assembly mechanisms of soil abundant and rare bacterial taxa under increasing pyrene stresses. Front Microbiol. 2021;12(689762):1–14.

    Google Scholar 

  49. Hong C, Si Y, Xing Y, Li Y. Illumina MiSeq sequencing investigation on the contrasting soil bacterial community structures in different iron mining areas. Environ Sci Pollut Res. 2015;22(14):10788–99.

    Article  CAS  Google Scholar 

  50. Boros-Lajszner E, Wyszkowska J, Borowik A, Kucharski J. The response of the soil microbiome to contamination with cadmium, cobalt and nickel in soil sown with Brassica napus. Minerals. 2021;11(5):1–16.

    Article  Google Scholar 

  51. Flores C, Catita JAM, Lage OM. Assessment of planctomycetes cell viability after pollutants exposure. Antonie Van Leeuwenhoek. 2014;106(2):399–411.

    Article  CAS  PubMed  Google Scholar 

  52. Syranidou E, Thijs S, Avramidou M, Weyens N, Venieri D, Pintelon I, et al. Responses of the endophytic bacterial communities of Juncus acutus to pollution with metals, emerging organic pollutants and to bioaugmentation with indigenous strains. Front Plant Sci. 2018;15:459–83.

    Google Scholar 

  53. Brescia F, Sillo F, Franchi E, Pietrini I, Montesano V, Marino G, et al. The ‘microbiome counterattack’: insights on the soil and root-associated microbiome in diverse chickpea and lentil genotypes after an erratic rainfall event. Environ Microbiol Rep. 2023;15(6):459–83.

    Article  PubMed  PubMed Central  Google Scholar 

  54. Văcar CL, Covaci E, Chakraborty S, Li B, Weindorf DC, Frențiu T, et al. Heavy metal-resistant filamentous fungi as potential mercury bioremediators. J Fungi. 2021;7(5):1–21.

    Article  Google Scholar 

  55. Marx DH, Bryan WC. Pure culture synthesis of ectomycorrhizae by Thelepltora tevvestvis and Pisolithus Tinctorius on different conifer hosts. Can J Bot. 1970;48:639–43.

    Article  Google Scholar 

  56. Borovička J, Braeuer S, Walenta M, Hršelová H, Leonhardt T, Sácký J, et al. A new mushroom hyperaccumulator: cadmium and arsenic in the ectomycorrhizal basidiomycete Thelephora penicillata. Sci Total Environ. 2022;826:1–10.

    Article  Google Scholar 

  57. Borovička J, Sácký J, Kaňa A, Walenta M, Ackerman L, Braeuer S, et al. Cadmium in the hyperaccumulating mushroom Thelephora penicillata: intracellular speciation and isotopic composition. Sci Total Environ. 2023;855:1–9.

    Article  Google Scholar 

  58. Stroheker S, Dubach V, Sieber TN. Competitiveness of endophytic Phialocephala fortinii s.l. – acephala applanata strains in Norway spruce roots. Fungal Biol. 2018;122(5):345–52.

    Article  PubMed  Google Scholar 

  59. Haug I, Weiß M, Homeier J, Oberwinkler F, Kottke I. Russulaceae and Thelephoraceae form ectomycorrhizas with members of the Nyctaginaceae (Caryophyllales) in the tropical mountain rain forest of southern Ecuador. New Phytol. 2005;165(3):923–36.

    Article  CAS  PubMed  Google Scholar 

  60. Schreiner PR, Koide RT. Antifungal compounds from the roots of mycotrophic and non-mycotrophic plant species. New Phytol. 1993;123(1):99–105.

    Article  CAS  Google Scholar 

  61. Glynou K, Nam B, Thines M, Maciá-Vicente JG. Facultative root-colonizing fungi dominate endophytic assemblages in roots of nonmycorrhizal Microthlaspi species. New Phytol. 2018;217(3):1190–202.

    Article  PubMed  Google Scholar 

  62. Wang L, Han M, Li X, Yu B, Wang H, Ginawi A, et al. Mechanisms of niche-neutrality balancing can drive the assembling of microbial community. Mol Ecol. 2021;30(6):1492–504.

    Article  PubMed  Google Scholar 

  63. Chase JM, Myers JA. Disentangling the importance of ecological niches from stochastic processes across scales. Philos Trans R Soc B Biol Sci. 2011;366(1576):2351–63.

    Article  Google Scholar 

  64. Burns AR, Stephens WZ, Stagaman K, Wong S, Rawls JF, Guillemin K, et al. Contribution of neutral processes to the assembly of gut microbial communities in the zebrafish over host development. ISME J. 2016;10(3):655–64.

    Article  CAS  PubMed  Google Scholar 

  65. Likar M, Grašič M, Gaberščik A. Contribution of neutral processes to the assembly of microbial communities on Phragmites australis leaf litter. ABS. 2023;66(2):1–10.

    Google Scholar 

  66. Li W, Hu X, Liu Q, Yin C. Soil fungi are more sensitive than bacteria to short-term plant interactions of Picea Asperata and Abies faxoniana. Eur J Soil Biol. 2021;106:1–10.

    Article  CAS  Google Scholar 

  67. Dennis PG, Rushton SP, Newsham KK, Lauducina VA, Ord VJ, Daniell TJ, et al. Soil fungal community composition does not alter along a latitudinal gradient through the maritime and sub-antarctic. Fungal Ecol. 2012;5(4):403–8.

    Article  Google Scholar 

  68. Zhang J, Zhang J, Yang L. A long-term effect of Larix monocultures on soil physicochemical properties and microbes in northeast China. Eur J Soil Biol. 2020;96:1–10.

    Article  Google Scholar 

  69. Li H, Weng B, Sen, Huang FY, Su JQ, Yang XR. pH regulates ammonia-oxidizing bacteria and archaea in paddy soils in Southern China. Appl Microbiol Biotechnol. 2015;99(14):6113–23.

    Article  CAS  PubMed  Google Scholar 

  70. Plaza C, Courtier-Murias D, Fernández JM, Polo A, Simpson AJ. Physical, chemical, and biochemical mechanisms of soil organic matter stabilization under conservation tillage systems: a central role for microbes and microbial by-products in C sequestration. Soil Biol Biochem. 2013;57:124–34.

    Article  CAS  Google Scholar 

  71. García-Orenes F, Roldán A, Morugán-Coronado A, Linares C, Cerdà A, Caravaca F. Organic fertilization in traditional mediterranean grapevine orchards mediates changes in soil microbial community structure and enhances soil fertility. Land Degrad Dev. 2016;27(6):1622–8.

    Article  Google Scholar 

  72. Shi S, Nuccio EE, Shi ZJ, He Z, Zhou J, Firestone MK. The interconnected rhizosphere: high network complexity dominates rhizosphere assemblages. Ecol Lett. 2016;19(8):926–36.

    Article  PubMed  Google Scholar 

  73. Ma B, Wang Y, Ye S, Liu S, Stirling E, Gilbert JA, et al. Earth microbial co-occurrence network reveals interconnection pattern across microbiomes. Microbiome. 2020;8(1):1–12.

    Article  CAS  Google Scholar 

  74. Conn HJ. A possible function of actinomycetes in soil. J Bacteriol. 1916;1(2):197–207.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  75. vanVilet PC, Hendrix PF. Role of fauna in soil physical processes. In: Abbot LK, Murphy DV, editors. Soil biological fertility. Dordrecht: Springer; 2007. pp. 61–80.

    Google Scholar 

  76. Fajardo C, Costa G, Nande M, Botías P, García-Cantalejo J, Martín M, Pb. Cd, and Zn soil contamination: monitoring functional and structural impacts on the microbiome. Appl Soil Ecol. 2019;135:56–64.

    Article  Google Scholar 

  77. Zhang J, Shi Q, Fan S, Zhang Y, Zhang M, Zhang J. Distinction between Cr and other heavy–metal–resistant bacteria involved in C/N cycling in contaminated soils of copper producing sites. J Hazard Mater. 2021;402:1–9.

    Article  Google Scholar 

  78. Fierer N, Bradford MA, Jackson RB. Toward an ecological classification of soil bacteria. Ecology. 2007;88(6):1354–64.

    Article  PubMed  Google Scholar 

  79. Pan X, Raaijmakers JM, Carrión VJ. Importance of Bacteroidetes in host–microbe interactions and ecosystem functioning. Trends Microbiol. 2023;31(9):959–71.

    Article  CAS  PubMed  Google Scholar 

  80. Larsbrink J, McKee LS. Bacteroidetes bacteria in the soil: glycan acquisition, enzyme secretion, and gliding motility. Adv Appl Microbiol. 2020;110:63–98.

    Article  CAS  PubMed  Google Scholar 

  81. Riley MA, Gordon DM. The ecological role of bacteriocins in bacterial competition. Trends Microbiol. 1999;7(3):129–33.

    Article  CAS  PubMed  Google Scholar 

  82. Matano LM, Coyne MJ, García-Bayona L, Comstock LE. Bacteroidetocins target the essential outer membrane protein BamA of Bacteroidales symbionts and pathogens. ASM. 2021;12(5):1–13.

    Google Scholar 

  83. Coyte KZ, Schluter J, Foster KR. The ecology of the microbiome: networks, competition, and stability. Science. 2015;350(6261):663–6.

    Article  CAS  PubMed  Google Scholar 

  84. Nishisaka CS, Ventura JP, Bais HP, Mendes R. Role of Bacillus subtilis exopolymeric genes in modulating rhizosphere microbiome assembly. Environ Microbiome. 2024;19(1):1–17.

    Article  Google Scholar 

  85. Nelson DW, Sommers LE. Total carbon, organic carbon, and organic matter. In: Sparks D, Page AL, Helmke PA, Loeppert RH, Soltanpour PN, Tabatabai MA, et al. editors. Methods of soil analysis part 3: chemical methods. Madison: Soil Science Society of America; 1996. pp. 961–1010.

    Google Scholar 

  86. Tamames J, Puente-Sánchez F. SqueezeMeta, a highly portable, fully automatic metagenomic analysis pipeline. Front Microbiol. 2019;9(3349):1–10.

    Google Scholar 

  87. Buchfink B, Xie C, Huson DH. Fast and sensitive protein alignment using DIAMOND. Nat Methods. 2015;12(1):59–60.

    Article  CAS  PubMed  Google Scholar 

  88. McMurdie PJ, Holmes S, Phyloseq. An R package for reproducible interactive analysis and graphics of microbiome census data. PLoS ONE. 2013;8(4):e61217.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  89. Likar M, Grašič M, Stres B, Regvar M, Gaberščik A. Original leaf colonisers shape fungal decomposer communities of Phragmites australis in intermittent habitats. J Fungi. 2022;8(3):1–14.

    Article  Google Scholar 

  90. Marasco R, Mosqueira MJ, Fusi M, Ramond JB, Merlino G, Booth JM, et al. Rhizosheath microbial community assembly of sympatric desert speargrasses is independent of the plant host. Microbiome. 2018;6(1):1–18.

    Article  Google Scholar 

  91. Hartman K, van der Heijden MGA, Wittwer RA, Banerjee S, Walser JC, Schlaeppi K. Cropping practices manipulate abundance patterns of root and soil microbiome members paving the way to smart farming. Microbiome. 2018;6(1):1–14.

    Google Scholar 

Download references

Acknowledgements

The work was supported by the COST association (Grant CA19116 “Trace Metal Metabolism in Plants – PLANTMETALS”).

Funding

This work was funded by the Slovenian Research and Innovation Agency (research core funding No. P1-0212 and project funding J4-3091) and the Young Researcher Scholarship to V.B.

Author information

Authors and Affiliations

  1. Biotechnical Faculty, University of Ljubljana, Jamnikarjeva 101, Ljubljana, SI-1000, Slovenia

    Valentina Bočaj, Paula Pongrac, Helena Grčman & Matevž Likar

  2. Jožef Stefan Institute, Jamova 39, Ljubljana, SI-1000, Slovenia

    Paula Pongrac

  3. National Institute of Chemistry, Hajdrihova 19, Ljubljana, SI-1000, Slovenia

    Martin Šala

Authors
  1. Valentina Bočaj
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Paula Pongrac
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. Helena Grčman
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. Martin Šala
    View author publications

    You can also search for this author in PubMed Google Scholar

  5. Matevž Likar
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

V. B. conducted the investigation, performed formal analyses, and wrote the main manuscript text.P.P.: conceptualized the study, conducted the lab work and formal analyses connected to the ICP-MS method, assisted with the writing, and reviewed and edited the manuscript.H.G.: conducted the lab work connected to soil analysis, and reviewed and edited the manuscript.M.Š.: conducted the lab work connected to the ICP-MS method.M.L.: conceived and designed the study, performed the formal analyses, and coordinated the work related to this manuscript.

Corresponding author

Correspondence to Matevž Likar.

Ethics declarations

Ethics approval and consent to participate

All necessary permissions and licences were obtained before the collection. A voucher specimen was deposited in the Herbarium of the University in Ljubljana under identification LJU10147538.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.

Supplementary Information

The online version contains supplementary material available at [here will be placed the link of the paper if it will be published].

Additional information

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Electronic supplementary material

Below is the link to the electronic supplementary material.

Supplementary Material 1

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Bočaj, V., Pongrac, P., Grčman, H. et al. Rhizobiome diversity of field-collected hyperaccumulating Noccaea sp.. BMC Plant Biol 24, 922 (2024). https://doi.org/10.1186/s12870-024-05605-4

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s12870-024-05605-4

Keywords

BMC Plant Biology

ISSN: 1471-2229

Contact us