Abstract
Aims
Compare bacterial communities in non-vegetated soils and in the rhizosphere of Ni-hyperaccumulating or Ni-excluding plants from four serpentine sites of the Iberian Peninsula.
Methods
Bacterial communities in non-vegetated soil and in rhizosphere of Alyssum serpyllifolium, and Dactylis glomerata, were analysed using denaturing gradient gel electrophoresis (DGGE) of 16S rRNA gene fragments of the total bacterial community and of Alphaproteobacteria, Actinobacteria and Streptomycetaceae.
Results
Rhizosphere bacterial communities of, either the hyperaccumulators or the excluder plants, were significantly different from communities in non-vegetated soils, especially for Alphaproteobacteria and Actinobacteria. The main differences between the rhizobacterial communities of the hyperaccumulator and of the excluder corresponded to Alphaproteobacteria profiles. Communities in non-vegetated soil were significantly influenced by intrinsic soil properties while in rhizosphere soil these factors were replaced by parameters related to nutrient availability. Members of the genera Blastococcus, Geodermatophilus, Modestobacter, Rhodococcus, Pseudonocardia, Devosia and Hyphomicrobium were detected in the rhizosphere of the Ni-hyperaccumulating plants.
Conclusions
Bacterial communities significantly differed between hyperaccumulator and excluder rhizosphere and non-vegetated soil. Actinobacteria of the Geodermatophilaceae family and Alphaproteobacteria of the Hyphomicrobiaceae family predominated in the rhizosphere profiles of the Ni-hyperaccumulators. Alphaproteobacteria could participate in the tolerance of Poaceae to trace elements in ultramafic soils.
Similar content being viewed by others
Explore related subjects
Discover the latest articles, news and stories from top researchers in related subjects.Avoid common mistakes on your manuscript.
Introduction
The soils derived from ultramafic rocks are commonly referred to as ultramafic or serpentine soils (Proctor 1999). These soils are characterised by elevated concentrations of Mg, Fe and trace elements (TE) such as Co, Cr and Ni, a low Ca/Mg ratio, low concentration of essential plant nutrients such as N, P and K and a strong tendency to drought (Brooks 1987; Whittaker 1954). These characteristics generate a challenging environment for the establishment of plants, which is one of the reasons why serpentine areas often support a high proportion of endemic species (Brooks 1987; Galey et al. 2017; van der Ent et al. 2013).
The ecology of plants inhabiting ultramafic areas has been well documented, with many plant species showing mechanisms of stress-tolerance. In 1981, Baker defined three strategies whereby plants could persist in environments with high levels of toxic metals and classified them as accumulators, excluders and indicators. Some of the plants following an accumulating strategy are able to concentrate extremely high amounts of metals in their shoots and are called hyperaccumulators (Brooks et al. 1977). Among hyperaccumulators, over 450 taxa are able to accumulate Ni (Pollard et al. 2014; van der Ent et al. 2013). The Iberian Peninsula hosts two serpentine-endemic and Ni-hyperaccumulating subspecies of Alyssum serpyllifolium Desf.: Alyssum serpyllifolium subsp. lusitanicum from Galicia (NW Spain) and Trás-os-Montes (NE Portugal), and Alyssum serpyllifolium subsp. malacitanum from Andalucía (S Spain) (López González 1975; Brooks et al. 1981; Menezes de Sequeira and Pinto da Silva 1992). Hyperaccumulating plants are considered good candidates for application in phytoextraction technologies such as the phytomining of soils or substrates naturally enriched or contaminated with metals. On the contrary, excluders are tolerant to high metal concentrations in soil but metals are excluded from uptake into aerial tissues.
The study of microbial communities in ultramafic soils has received less attention. Studies based on traditional culture-dependent methods generally indicate that these soils are characterised by low microbial density, biomass and activity, which has been attributed mainly to the high concentrations of TE (Acea and Carballas 1986; Lipman 1926; Pal et al. 2005). More recently, some authors using molecular analyses detected high microbial diversity in ultramafic soils (Bordez et al. 2016; Mengoni et al. 2004; Oline 2006; Thijs et al. 2017; Lopez et al. 2017). Recently Lopez et al. (2017) using a high-throughput sequencing technique found, in the rhizosphere of the hyperaccumulator Alyssum murale growing in ultramafic soils in Greece, a highly diverse bacterial community dominated by Chloroflexi. These authors also observed an important influence of Ni availability (which was positively correlated with the relative abundance of Alphaproteobacteria and Actinobacteria) on the composition of the bacterial communities. Ultramafic soils are particularly interesting for studying natural microbial adaptation to metal toxicity and the evolution of microbial communities in soils naturally enriched in TE, which constitutes a situation very different from that found in anthropogenically contaminated soils (Héry et al. 2003). Most studies assessing the microbiology of ultramafic soils are devoted to the analysis of bacterial communities in the rhizosphere of Ni-hyperaccumulating plants (Visioli et al. 2015).
The rhizosphere represents a complex microenvironment where plant roots and soil interact with microorganisms. Rhizosphere microorganisms play a vital role in the biogeochemical cycling of nutrients and in the speciation and bioavailability of metals for plants (Kuffner et al. 2008; Mendes et al. 2013; Muehe et al. 2015; Philippot et al. 2013). Likewise, plants exert a great influence on microorganisms through the excretion of root exudates (Philippot et al. 2013). These exudates are an important source of nutrients that will be used by soil bacteria for energy and biomass production (Haichar et al. 2008; Ma et al. 2016). Moreover, these exudates take part in the early colonisation of the roots by indigenous soil microorganisms, inducing chemotactic response of rhizosphere bacteria (Dennis et al. 2010; Haichar et al. 2008; Segura et al. 2009). Root exudates are considered the main factor affecting indigenous soil microorganisms and driving the establishment of specific microbial communities in the rhizosphere of different plant species (Berg and Smalla 2009).
Studies of the rhizosphere bacterial communities associated with Ni-hyperaccumulating plants have been mainly focused on the culturable bacterial community and aimed at the isolation and characterisation of bacterial strains with potential use as bioinoculants to improve metal phytoextraction (Abou-Shanab et al. 2003; Álvarez-López et al. 2016; Becerra-Castro et al. 2009; Benizri and Kidd 2018; Idris et al. 2004; Mengoni et al. 2001). Microbial communities associated with non-accumulating plants growing in ultramafic environments have received much less attention (Mengoni et al. 2010). A few authors have conducted comparative studies looking at the culturable bacterial communities associated with either Ni-hyperaccumulators or non-accumulating plants (Álvarez-López et al. 2016; Becerra-Castro et al. 2009; Delorme et al. 2001), while culture-independent techniques have not been used in this type of study. Becerra-Castro et al. (2009) and Álvarez-López et al. (2016) found higher densities of Ni-tolerant bacteria in the rhizosphere of some populations of the Ni-hyperaccumulator Alyssum serpyllifolium than in the rhizosphere of Dactylis glomerata, a metal excluding plant, or non-vegetated soil from the same sites. The increase in densities of metal-tolerant bacteria was related to a higher concentration of bioavailable metal in the rhizosphere of the hyperaccumulators (Becerra-Castro et al. 2009). These authors also observed some differences in the taxa isolated from either the rhizosphere of the hyperaccumulating or excluding plants (Álvarez-López et al. 2016). The rhizosphere of the metal hyperaccumulating Noccaea caerulescens (syn. Thlaspi caerulescens) growing in Zn and Cd contaminated soil was also found to host a higher ratio of metal-resistant bacteria than the non-hyperaccumulating Trifolium pratense growing in the same soil (Delorme et al. 2001).
The aim of this study was to analyse the bacterial community present in the rhizosphere of two Ni-hyperaccumulating subspecies of Alyssum serpyllifolium Desf. (Alyssum serpyllifolium subsp. lusitanicum and Alyssum serpyllifolium subsp. malacitanum) and the metal-excluder plant, Dactylis glomerata L., growing in four ultramafic areas in the Iberian Peninsula. The bacterial communities were studied using denaturing gradient gel electrophoresis (DGGE) of PCR-amplified 16S rRNA gene fragments obtained with primers designed for Bacteria as well as with group-specific primers. Additionally, multivariate statistical analyses were used to study the relationship between soil properties and the composition of the bacterial communities.
Materials and methods
Study sites and sample collection
Samples were collected in four serpentine areas of the Iberian Peninsula: Barazón (L), in Galicia, NW of Spain (42°51′09″N; 08°01′15″W), Morais (M) and Samil (S), in Trás-os-Montes, NE of Portugal (41°31′21″N; 06°49′20″W and 41°46′48″N; 6°44′47″W, respectively) and Sierra Bermeja (SB) in Andalucía, South of Spain (36°28′48″N; 05°11′52″W). In these serpentine areas, two Ni-hyperaccumulating subspecies of Alyssum serpyllifolium Desf., endemic to the Iberian Peninsula are found: A. serpyllifolium subsp. lusitanicum Dudley and P. Silva (commonly referred as A. pintodasilvae) in L, M and S sites, and A. serpyllifolium subsp. malacitanum Rivas Goday (A. malacitanum) in SB site (Brooks et al. 1981; Gómez-Zotano et al. 2014; Menezes de Sequeira 1969). The whole plant and root system (including root ball) of 5–7 individuals of the four mentioned Alyssum populations were collected. In L, M and S sites 5–7 individuals of the Ni-excluder, Dactylis glomerata subsp. hispanica (Roth) Nyman were also collected. The rhizosphere soil was separated by gently crushing the root ball and shaking the root system. Tightly held soil (<3 mm from the root surface) was considered as rhizosphere soil. In addition, surface soil samples (0–10 cm) were collected at each site from bare patches at approximately 1 m distance from the sampled Alyssum individuals (non-vegetated soil). Aliquots (0.5 g) of the rhizosphere and bulk soil samples were stored at −60 °C until DNA extraction. The analysis of bacterial communities was carried out in the rhizosphere of 4 individuals of each population as well as in 4 replicates of non-vegetated soil from the different serpentine sites. The remaining samples were air-dried and used for physico-chemical analyses, which were also carried out in four replicates as described by Álvarez-López et al. (2016). The main physico-chemical characteristics of the soils at each site are listed in the Table SI 1.
The nomenclature of samples in this study included a first letter indicating the mentioned sampling sites (L, Barazón; M, Morais; S, Samil; SB, Sierra Bermeja), followed by AR, GR or NV, denoting the rhizosphere of Alyssum, the rhizosphere of Dactylis or non-vegetated soil, respectively.
DNA extraction and PCR amplification of 16S rRNA gene fragments for DGGE
Total soil DNA was extracted from 0.5 g of soil using the PowerSoil® DNA Isolation Kit (MoBio Laboratories Inc., California) following the manufacturer’s instructions. DNA quality was checked in a 0.8% agarose gel.
The amplification of 16S rRNA gene fragments of Alphaproteobacteria, Actinobacteria and Streptomycetaceae was carried out using a (semi)nested PCR approach. A non-nested approach was used to amplify 16S rRNA gene fragments of the total bacterial community. PCR primers and conditions used are shown in Table SI 2 and PCR reactions were carried out as described by Touceda-González et al. (2017).
DGGE analysis of amplified 16S rRNA gene fragments
A phorU2 apparatus was used to perform the DGGE analysis (Ingeny, Goes, The Netherlands) of amplified 16S rRNA gene fragments using a double (acrylamide and denaturant) gradient. The gradient was composed of 46.5 to 65.0% of denaturant (100% denaturant corresponds to 7 M urea and 40% (v/v) formamide) and 6.2 to 9.0% acrylamide (Gomes et al. 2005). The amount of PCR product used in DGGE was 5 μl, for group–specific fragments, or 7 μl, for total bacterial community fragments. The gels were run in 1× TAE buffer at a constant voltage of 140 V for 17 h at 58 °C, and the gel was stained following a conventional silver staining procedure. After electrophoresis the gels were air-dried and scanned (Epson Perfection V750 Pro, Japan). A marker composed of GC-clamped 16S rRNA gene fragments (positions 984 to 1378) of 7 previously isolated bacterial strains with different electrophoretic mobility was loaded in two lanes on each gel.
Analysis of DGGE fingerprints
The DGGE profiles were analysed using the GelCompar II program, version 6.0 (Applied Maths, Ghent, Belgium). The marker loaded on every gel was used as a reference to convert and normalise the gel images. The pairwise similarities of lanes were calculated for each gel using the Pearson correlation coefficient and the resulting similarity matrices were used for cluster analysis by the unweighted pair-group method using average linkages (UPGMA). The statistical significance of the differences between the microbial community present in the rhizosphere of Alyssum or Dactylis and in the non-vegetated soil was analysed with a permutation test (Kropf et al. 2004) using the Pearson correlation coefficients similarity matrices. Canonical Correspondence Analysis (CCA), carried out with CANOCO 5 software (Microcomputer Power, USA), was used to relate DGGE fingerprinting information to environmental data (physico-chemical soil properties). The matrices containing DGGE information consisted of relative band intensities calculated by dividing individual band intensity by the sum of all the band intensities in the analysis. Explanatory variables were transformed to a normal distribution by scaling the data obtaining a mean of 0 and a standard deviation of 1. The explanatory variables showing a significant influence on the composition of the bacterial communities (P < 0.05) were chosen using manual forward selection (i.e. model building was initiated including the variable that is most significant in the initial analysis, and continued to add variables until the model was not improved by the incorporation of a new variable). The statistical significance of the canonical axes was tested with a Monte Carlo permutation test (n = 999). The DGGE bands which were best explained by the model (FitE values >75%) were selected.
Excision and sequencing of DGGE fragments
Some of the DGGE bands, which were sample specific or more intense in one sample compared to the others, were excised and sequenced. Bands were cut from the gels by using a sterile scalpel, and immediately placed in a centrifuge tube, broken into small pieces with a sterile pipet tip and incubated overnight at 4 °C in 50 μl of sterile Mili-q water for the elution of DNA. The tubes containing the DGGE bands were centrifuged at 10,000 g for 10 min and the supernatant was carefully recovered. The eluted DNA was then re-amplified using primers F984GC and R1378 as previously described. PCR reamplified fragments were ligated into the pCR™4-TOPO® vector (ThermoFischer) and used for the transformation of One Shot® Top10-competent Escherichia coli cells (ThermoFischer). Blue-white screening of transformants was done on LB agar plates containing 50 mg mL−1 of ampicillin and top spread with 40 μL of 40 mg mL−1 of X-Gal (5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside). Clones were screened for inserts of the correct size by PCR with appropriate primers included in the cloning kit. Three clones from each excised band were sequenced. The obtained sequences were compared to those in the EMBL database by using FASTA at the European Bioinformatics Institute and with those of the GenBank by using nucleotide BLAST to identify closely related gene sequences.
Results
Bacterial community analysis
All PCR amplified fragments of 16S rRNA gene of Bacteria (total bacterial community, TC) Alphaproteobacteria, Actinobacteria and Streptomycetaceae generated products which were adequate for analysis by DGGE.
Complex band patterns were obtained for all sites and bacterial groups analysed. In general, there was a good reproducibility between biological replicates both in the rhizosphere and in the non-vegetated soil samples. The UPGMA dendrograms (based on the Pearson similarity coefficients) showed that rhizosphere DGGE profiles of the hyperaccumulator (AR) could be distinguished from those of the excluder rhizosphere (GR) or of the non-vegetated soil (NV) (Fig. 1).
The dendrogram obtained from the similarity matrix of DGGE profiles for the total bacterial community showed a cluster including all samples from the Portuguese sites (Fig. 1a). Within this cluster, the rhizosphere samples of Dactylis and Alyssum (except for two replicates) were separated into two different groups and the non-vegetated soils, SNV and MNV, also formed two additional separate groups (Fig. 1a). In contrast, samples from Barazón and Sierra Bermeja did not form separate groups. Instead, LAR replicates clustered together and showed low similarity with other samples. SBAR samples also grouped together and surprisingly appeared associated to LNV (Fig. 1a).
In the dendrogram of Alphaproteobacteria, the rhizosphere of all Dactylis populations clustered within the same clade (Fig. 1b). On the contrary, the rhizosphere of Alyssum grouped in two distant clusters, the first including samples from Sierra Bermeja and most of the replicates of Samil and the second containing samples from Barazón and all other replicates of the Portuguese populations. Alphaproteobacteria profiles of the non-vegetated soil from Barazón and Sierra Bermeja formed two distant groups with a high similarity between replicates but the non-vegetated soil from Samil and Morais showed more heterogeneous profiles (Fig. 1b).
In the dendrogram of Actinobacteria, the samples from the rhizosphere of Alyssum populations formed four distant clades (Fig. 1c). In general, the Alyssum profiles were also distantly related to the patterns found in the corresponding non-vegetated soils. The profiles of the rhizosphere of Dactylis populations also constituted separate clusters. In Barazón and Morais the Actinobacteria associated with Dactylis were closer to those of the non-vegetated soil than to the Alyssum community (Fig. 1c).
In the dendrogram of Streptomycetaceae, the samples from different populations tended to group separately. The profiles from Sierra Bermeja formed a cluster (Fig. 1d) and within this the profiles of Alyssum rhizosphere (SBAR) clustered separately from those of non-vegetated soil (SBNV). Most of the samples from Portugal, except for the non-vegetated soil from Samil and two rhizosphere replicates from Morais formed a big clade. Within this clade, the samples from Samil were grouped and the rhizosphere of Alyssum clustered separately from that of Dactylis. The samples from Morais, also tended to group together but the profiles were more heterogeneous. Most samples from Barazón also formed a single cluster and these were more related to the Portuguese populations than to the Sierra Bermeja population. Within this clade the non-vegetated soil profiles (LNV) clustered separately from the rhizosphere profiles.
On a site-by-site level, most of the dendrograms of Bacteria DGGE profiles revealed a remarkable separation in community structure between Alyssum and Dactylis and that of the non-vegetated soil. Based on the permutation tests, differences between the DGGE fingerprints of the rhizosphere, either Alyssum or Dactylis, and of the non-vegetated soil, were statistically significant, except in the case of some of the Streptomycetaceae profiles (P < 0.05; Table 1). Generally, the most pronounced dissimilarities corresponded to the profiles of total bacterial community, although the calculated d-value for the comparison between Alyssum rhizosphere and the non-vegetated soil was maximal for the Alphaproteobacterial profiles in Barazón and for the Actinobacterial profiles in Samil and Sierra Bermeja (P < 0.05; Table 1). The fingerprints of total bacterial community, Alphaproteobacteria and Actinobacteria of the Alyssum rhizosphere were also statistically different from those of the Dactylis rhizosphere. The dissimilarities reached maxima in the case of Alphaproteobacterial profiles.
The analysis of DGGE profiles from Alyssum rhizosphere growing in Barazón, Morais, Samil and Sierra Bermeja revealed that all four rhizosphere soils host significantly different bacterial communities (P < 0.05; Table 1), and this was generally the case for all the studied bacterial groups (Fig. SI 1). The lower calculated dissimilarities corresponded to the differences between the two Portuguese populations. The dissimilarities tended to be higher in the case of the Streptomycetaceae profiles. In the Alphaproteobacteria profiles there was a noteworthy high similarity between SBAR replicates.
Relationship between environmental variables and the bacterial community DGGE patterns
CCA was used to investigate the correlation between environmental variables and the bacterial community data from DGGE patterns.
The data corresponding to the rhizosphere of the two plant species and to the non-vegetated soils were analysed separately and the explanatory variables obtained in the CCAs were compared (Figs. 2 and SI 2). In the CCA of DGGE band patterns of the different bacterial groups the variance explained by axis 1 and 2 was generally around 30%, although in the Alphaproteobacteria CCA of the Alyssum rhizosphere (Fig. 2a) and in the Actinobacteria CCA of Alyssum and Dactylis rhizosphere (Fig. 2d, e), only one axis was extracted by the analysis and the variance explained was lower (between 13 and 22%). The environmental variables contributed to explaining the variation in the community of all studied bacterial groups (the species-environment correlations for axes 1 and 2 were generally >0.885 and > 0.812, respectively).
In general, the CCA biplots defined by axes 1 and 2 (Figs. 2c, f and Figs. SI 2 c and SI 2 f) showed a separation of the samples of non-vegetated soils from the four serpentine sites studied. A similar grouping was observed for the samples of the Dactylis rhizosphere except in the CCA biplot of the Actinobacterial band patterns (Figs. 2b, e and Figs. SI 2 b and SI 2 e). In the case of Alyssum rhizosphere, the samples from Barazón were generally separated from all other Alyssum samples, although in the CCA of the total bacterial community samples from Sierra Bermeja appeared together with those of Barazón (Figs. 2a, d and Figs. SI 2 a and SI 2 d).
The variables which contributed most to explaining variations in the bacterial community depended on the type of soil and on the bacterial group. In the case of the total community patterns of the non-vegetated soil (Fig. SI 2 c), the variables with the highest intraset correlations were exchangeable Mg and K and CEC. Nevertheless, exchangeable K was absent in the CCA of samples of Alyssum rhizosphere (Fig. SI 2 a) and CEC absent in the CCA of Dactylis rhizosphere (Fig. SI 2 b). Additionally, in the latter case, total Ni concentration also showed a high loading factor on axis 2.
The environmental variables which mostly influenced Alphaproteobacteria, Actinobacteria and Streptomycetaceae community composition of the non-vegetated soil were total organic C, total Ni concentration and available Mg (Figs. 2c, f and SI 2 f). In the CCA of Streptomycetaceae CEC also showed high loading factors on both axes 1 and 2 (Fig. SI 2 f). However, the variables explaining the variation in these bacterial groups in the rhizosphere of Alyssum and Dactylis were different. In the rhizosphere of Alyssum only CEC showed a high intraset correlation with axis 1 of the CCA of Alphaproteobacteria and Actinobacteria, while in the Streptomycetaceae CCA, CEC presented a high loading factor on axis 1 and EDTA-extractable Co on both axes 1 and 2 (Figs. 2a, d and SI 2 d). In the rhizosphere of Dactylis, EDTA-extractable Ni and exchangeable Mg contributed to explaining the variation in Alphaproteobacteria, exchangeable K affected the variation of Actinobacteria, while in the Streptomycetaceae CCA the variables with highest intraset correlations were EDTA-exchangeable Ni, Co and Cr (Figs. 2b, e and SI 2 e).
Separate CCA analyses were also carried out for each bacterial group combining data from the non-vegetated soil with either Alyssum or Dactylis rhizosphere (Fig. SI 3). The variance explained by axes 1 and 2 was generally low (15–18%). In these analyses the environmental variables more related with the bacterial composition were different in the case of Alyssum or Dactylis rhizosphere. In the CCA biplots of Alphaproteobacteria, Actinobacteria and Streptomycetaceae the samples of non-vegetated soil were separated from those of Dactylis rhizosphere (Fig. SI 3a to 3f). The variables that contributed most to this separation were CEC and available K. In the case of Alyssum rhizosphere, the separation from non-vegetated samples was not consistent and available K was absent among the significant variables (Fig. SI 3a, 3c, 3e and 3 g).
Sequence analysis of dominant DGGE bands
Some prominent and characteristic bands observed in the DGGE profiles of the rhizospheres or the non-vegetated soil were excised and processed for their nucleotide sequence analysis (Fig. 3).
In the DGGE gel of total bacterial community few of the excised bands could be sequenced (Fig. 3a). Most of the sequences obtained were classified within phylum Actinobacteria, although none of them could be affiliated to specific organisms (Table SI 3).
In the case of Alphaproteobacteria and Actinobacteria, in general, the sequence of clones obtained from one band showed a high similarity and were identified as belonging to the same or phylogenetically close organisms (Tables 2 and 3).
Several bands characteristic of the Alyssum rhizosphere were cut from the Alphaproteobacteria DGGE gel (Fig. 3b). One additional prominent band present in SBNV was also analysed. The sequence of most of the prominent bands analysed showed high similarity with the 16S rRNA gene sequence of Devosia spp. and Hyphomicrobium spp. belonging to Hyphomicrobiaceae family (Table 2).
In the Actinobacteria DGGE gel, prominent bands characteristic of the rhizosphere of the different Alyssum populations, (Fig. 3c) as well as some bands from the profiles of non-vegetated soil of Samil and Sierra Bermeja and of the rhizosphere of Dactylis from Barazón and Morais, were analysed. The sequences of bands obtained from the Alyssum rhizosphere were highly similar to those of Modestobacter, Blastococcus or Geodermatophilus genera of the family Geodermatophilaceae (Table 3). Bands with sequences closer to that of Rhodococcus erythropolis, Mycobacterium or Pseudonocardia were also prominent in the Alyssum rhizosphere profiles. Intense bands identified as from Mycobacterium and Pseudonocardia were detected in the profiles of Dactylis rhizosphere and of non-vegetated soil, although their electrophoretic mobility was different than those detected in the Alyssum rhizosphere profiles. Other prominent bands in the profiles of non-vegetated soil were identified as derived from Tsukamurella sp. and Arthrobacter sp. (Table 3).
Discussion
In this study, DGGE analysis of 16S rRNA gene fragments has been applied to study the bacterial communities in non-vegetated soil and in the rhizosphere of Ni-hyperaccumulating (subspecies of Alyssum serpyllifolium) and the excluder (Dactylis glomerata) growing in four serpentine areas in the Iberian Peninsula. In the four studied sites, the bacterial communities in the rhizosphere soils significantly differed from the communities in non-vegetated soils, and this was observed at different taxonomic levels (Bacteria, Alphaproteobacteria, Actinobacteria and Streptomycetaceae).
Several studies have shown prominent differences between the microbial communities in non-vegetated and rhizosphere soils (Costa et al. 2006; DeAngelis et al. 2008; Gomes et al. 2001; Smalla et al. 2001). The rhizosphere is a stable and favourable environment that favours the settlement of soil microorganisms and acts selectively on the composition and structure of microbial communities (Steer and Harris 2000). Numerous studies have demonstrated increased microbial biomass and activity in the rhizosphere, which has mainly been related to the release of rhizodeposits by plant roots, principally as root exudates (Berg and Smalla 2009; Brimecombe et al. 2001; Hartmann et al. 2009). Root exudates act as signalling molecules and are a major source of available carbon for microbial proliferation (Brimecombe et al. 2001; Hartmann et al. 2009). Many soil microbes are carbon limited, whereby a quick response to root-induced changes in the rhizosphere soil is expected (DeAngelis et al. 2008). In our study, the organic matter content and dissolved organic C tended to be higher in the rhizosphere than in the non-vegetated soil. In addition, in low nutrient soils (such as serpentine soils) a stronger dependence of the root microbiome on plant exudates is expected (Dakora and Phillips 2002; Marschner et al. 2004).
Here, the rhizosphere effect was similar or more pronounced for Alphaproteobacteria and/or Actinobacteria than for the total bacterial community. Several authors found that the intensity of the rhizosphere effect depends on the bacterial group under investigation (Costa et al. 2006; Garbeva et al. 2004; Gomes et al. 2001). Gomes et al. (2001) found a strong seasonal population shift in the rhizosphere community of maize cultivars in the DGGE patterns of total bacteria, Alpha- and Beta-proteobacteria, and to a lesser extent for Actinobacteria. In this study, we focused on Actinobacteria and Proteobacteria bacterial groups based on the results obtained in previous culture-dependent studies which found that the culturable rhizosphere bacterial communities of the Ni-hyperaccumulating Alyssum in the Iberian Peninsula were dominated by members of these two phyla (Álvarez-López et al. 2016; Becerra-Castro et al. 2009; Becerra-Castro et al. 2011). Members of Proteobacteria and Actinobacteria are considered as abundant inhabitants of the rhizosphere (Philippot et al. 2013). The Proteobacteria phylum includes numerous fast-growing bacteria, known as r strategists, which have the capacity to use a wide range of root exudates as carbon substrate (Fierer et al. 2007; Philippot et al. 2013) as well as numerous plant symbiotic and plant growth promoting bacteria able to fix atmospheric nitrogen (Gomes et al. 2001; McCaig et al. 1999).
The UPGMA dendrograms and the permutation test, showed significant differences between the bacterial communities associated with the hyperaccumulator and excluder plants in the serpentine sites analysed. Several studies have highlighted the influence of different plant species on the composition and structure of rhizosphere bacterial communities (Costa et al. 2006; DeAngelis et al. 2008; Dias et al. 2012; Haichar et al. 2008; Hartmann et al. 2009; Smalla et al. 2001; Steer and Harris 2000). Smalla et al. (2001), detected plant species-specific bacterial communities in the rhizosphere of three important agricultural plant species, strawberry, oilseed rape and potato. Haichar et al. (2008) worked with plant species belonging to the same families as Alyssum (Brassicaceae) and Dactylis (Poaceae) and found that the differences between the rhizosphere communities of rape (Brassicaceae) and wheat (Poaceae) were more pronounced in the case of bacteria using root exudates than when analysing other bacteria present in the rhizosphere.
The hyperaccumulating Alyssum and the excluder Dactylis have developed opposing mechanisms to cope with high concentrations of TE which are naturally present in the ultramafic soils where they proliferate. Hyperaccumulators accumulate high concentrations of TE in their shoots, while excluders limit the absorption and transfer of metals to the aboveground biomass (Baker 1981). Several authors have suggested that root exudates, especially chelating organic acids, may play an important role in Ni acquisition by hyperaccumulators (Benizri and Kidd 2018; Wenzel et al. 2003). Poaceae are known for their capacity to produce compounds such phytosiderophores, which have a high affinity for iron and are also able to chelate other micronutrients. Some authors found differences in the composition and chelating properties of organic compounds in the rhizosphere of hyperaccumulators and excluders (Cattani et al. 2009; Wenzel et al. 2003). In the sites studied here, the concentration of dissolved organic carbon in the rhizosphere of Dactylis was higher (in Barazón and Morais) or similar (in Samil) than in the rhizosphere of Alyssum. However, we did not study the exudate composition of these two plant species. Changes induced by plants in the soil surrounding the roots are also affected by the root system architecture (Hinsinger and Courchesne 2007; Schwartz et al. 1999). Differences in root size and morphology between Alyssum and Dactylis may also have an impact on the rhizosphere bacterial community.
The dissimilarities between the rhizosphere communities of the two plants reached their maximum in the case of Alphaproteobacteria. Moreover, Alphaproteobacterial profiles of the rhizosphere of different Dactylis populations showed a high similarity and clustered together separately from the other soils. Gomes et al. (2010) observed that the soil amendment with non-contaminated and Cd- and Zn-contaminated sludges induced a more pronounced alteration of the Alphaproteobacteria community in arable soils than in soils covered by Poaceae (permanent grass ley). The differences observed were related to the potential plasmid-mediated transfer of heavy metal resistance within Alphaproteobacteria (Sandaa et al. 1999, 2001). Lopez et al. (2017) also found a positive correlation between Ni availability and the relative abundance of some Alphaproteobacteria such as members of the family Hyphomonadaceae, in ultramafic soils in Greece. Alphaproteobacteria may play a role in the mechanisms of exclusion or contribute to the tolerance of Poaceae to the high concentrations of TE in ultramafic soils.
The CCA analysis indicated that the main environmental factors influencing the bacterial community in the non-vegetated soil differed from those affecting the communities in the rhizosphere of Alyssum and Dactylis. In the non-vegetated soils, the Alphaproteobacteria, Actinobacteria and Streptomycetaceae communities were affected by available Mg and soil characteristics such as the total organic matter content and total Ni concentration. However, in the rhizosphere soils the influence of total Ni and organic matter content was replaced by other parameters related to nutrient availability (available K or Mg in the rhizosphere of Dactylis, and CEC in the rhizosphere of Alyssum). In the Dactylis rhizosphere, but not in the Alyssum rhizosphere, the available concentration (extractable with EDTA) of TE also contributed towards shaping the communities of Alphaproteobacteria and Streptomycetaceae. The results suggest a potential role of the rhizosphere bacterial community on plant nutrition and, in the case of the excluder, in TE tolerance. Despite similar levels of available K in the rhizosphere of both Dactylis and Alyssum, this variable contributed towards shaping the bacterial communities associated with the excluder but not with the hyperaccumulator. This result could be related to the contrasting strategies of these plants and their associated microorganisms for scavenging nutrients in ultramafic soils.
Some dominant bands were excised from DGGE gels with the aim of identifying members of the bacterial community characteristic of the different samples analysed. We observed that some bands, particularly those from the total community profiles, were constituted by fragments from organisms phylogenetically very distant which indicates that a large diversity might be hidden behind a single DGGE band. Similar observations were made by several authors (Costa et al. 2006; Gomes et al. 2001; Schmalenberger and Tebbe 2003; Smalla et al. 2001). Furthermore, bands excised from different positions in the gels were found to be associated with organisms of the same genus. Kušar and Avguštin (2012) obtained similar results when optimising the DGGE band identification method.
The DGGEs targeting different bacterial groups allowed for the detection of organisms that were not found in the total community analysis, enhancing the level of resolution of the PCR-DGGE technique. The sequences of bands characteristic of Alyssum rhizosphere excised from the actinobacterial DGGE gels were found to be originated from members of the genera Blastococcus sp., Geodermatophilus sp. and Modestobacter sp. (all three belonging to the family Geodermatophilaceae), Mycobacterium sp., Rhodococcus sp. and Pseudonocardia sp. The bacteria affiliated to the family Geodermatophilaceae have the ability to colonise poor substrates, such as rocks and extreme soils of hot and cold deserts and present modest growth requirements (Normand et al. 2014). Members of Geodermatophilaceae possess enzymatic tools to adapt to extreme environments. Essoussi et al. (2010) observed that the esterase activities in some members of this family, showed a high flexibility under high temperature, alkaline pH and high cationic concentration. Gtari et al. (2012) found a higher resistance to TE in Blastococcus than in Geodermatophilus obscurus or Modestobacter multiseptatus. The bands with a sequence similar to that of Pseudonocardia sp. were excised from both the non-vegetated ultramafic soil and the rhizosphere of Alyssum, but band intensity was higher in the rhizosphere profiles. Bacteria from this genus have been isolated and commonly detected in association with plants (Franco and Labeda 2014). Álvarez-López et al. (2016) also isolated strains of Lentzea sp., from the rhizosphere of the Alyssum. In the two Portuguese populations, a band characteristic of the Alyssum rhizosphere showed a sequence similar to that of Rhodococcus erythropolis. Members of the genus Rhodococcus are widely distributed in soils, have a saprophytic lifestyle and they are known to have a role in organic matter turnover and in the degradation of xenobiotics (Goodfellow 2014). Rhodococcus strains represented up to 10% of the isolates obtained (Álvarez-López et al. 2016; Becerra-Castro et al. 2011) from the rhizosphere of A. serpyllifolium subsp. lusitanicum and subsp. malacitanum growing in the Iberian Peninsula. Rhodococcus sp. were also found in the rhizosphere of the hyperaccumulator, Noccaea goesingensis (Idris et al. 2004).
Lopez et al. (2017), when analysing the rhizosphere bacterial community of the Ni-hyperaccumulator Alyssum murale growing in serpentine soils in Greece, found that the relative abundance of Actinobacteria, particularly of members of the family Gaiellaceae, was positively correlated with Ni availability. Gremion et al. (2003) also observed that Actinobacteria, in particular members of Rubrobacteria genus, are a dominant part of the metabolically active population in the rhizosphere of the Cd and Zn hyperaccumulator Noccaea caerulescens.
In this study, members of the Hyphomicrobiaceae family (Alphaproteobacteria) were detected in the rhizosphere of Alyssum, in particular members of the genera Devosia and Hyphomicrobium. One Devosia strain was isolated from the rhizosphere of A. serpyllifolium subsp. lusitanicum by Becerra-Castro et al. (2011). Genes for both nitrogen fixing and nodulation were found in Devosia neptuniae isolated from an aquatic leguminous plant (Rivas et al. 2002, 2003). Other Devosia strains were cultivated from root nodules; however, the role of these organisms in N2 fixation has not been demonstrated. Some Hyphomicrobiaceae bacteria such as members of Hyphomicrobium are characterised by the presence of prosthecate, monopolar or bipolar semi-rigid filamentous appendages that confer them the ability to survive in nutrient poor habitats as the prosthecate increase the cells surface area to increase uptake of nutrients (Oren and Xu 2014). Hyphomicrobiaceae bacteria, including members of the genera Devosia and Hyphomicrobium were among the most represented Alphaproteobacteria in the rhizosphere of Ni-hyperaccumulator Alyssum murale (Lopez et al. 2017). Mengoni et al. (2004) also detected some Alphaproteobacteria characteristic of the rhizosphere of the Ni-hyperaccumulator Alyssum bertolonii, although the method used (T-RFLP of 16S rRNA gene fragments) did not offer detailed taxonomic information.
Overall, in the four ultramafic sites studied the rhizosphere bacterial communities were significantly different from the communities in non-vegetated soils and this was observed at different taxonomic levels. The rhizobacterial communities associated with the hyperaccumulators and excluder plants also differed significantly. The sequences of bands characteristic of Alyssum rhizosphere indicated that Actinobacteria of the family Geodermatophilaceae (genera Blastococcus, Geodermatophilus, or Modestobacter) and of genera Rhodococcus and Pseudonocardia, as well as Alphaproteobacteria of genera Devosia and Hyphomicrobium, are likely to be relevant members of the rhizobacterial community associated with this hyperaccumulator plant. Regarding the Ni-excluder Dactylis, the clustering of Alphaproteobacteria rhizospheric profiles of different populations and the high dissimilarity found with Alphaproteobacteria in the rhizosphere of the hyperaccumulating plants suggest that members of this class could play a role in the mechanisms of exclusion or contribute to the tolerance of Poaceae to the high concentrations of TE in metal enriched soils.
References
Abou-Shanab RA, Angle JS, Delorme TA, Chaney RL, van Berkum P, Moawad H, Ghanem K, Ghozlan HA (2003) Rhizobacterial effects on nickel extraction from soil and uptake by Alyssum murale. New Phytol 158:219–224
Acea MJ, Carballas T (1986) Relationships among microbial groups in various humid zone soils, and the factors controlling their distribution. Agrochimica 34:1–14
Álvarez-López V, Prieto-Fernández Á, Becerra-Castro C, Monterroso C, Kidd PS (2016) Rhizobacterial communities associated with the flora of three serpentine outcrops of the Iberian Peninsula. Plant Soil 403:233–252
Baker AJM (1981) Accumulators and excluders - strategies in the response of plants to heavy-metals. J Plant Nutr 3:643–654
Becerra-Castro C, Monterroso C, Garcia-Leston M, Prieto-Fernandez A, Acea MJ, Kidd PS (2009) Rhizosphere microbial densities and trace metal tolerance of the nickel hyperaccumulator Alyssum serpyllifolium subsp. lusitanicum. International Journal of Phytoremediation 11:525–541
Becerra-Castro C, Prieto-Fernández A, Álvarez-Lopez V, Monterroso C, Cabello-Conejo MI, Acea MJ, Kidd PS (2011) Nickel solubilizing capacity and characterization of rhizobacteria isolated from hyperaccumulating and non-hyperaccumulating subspecies of Alyssum serpyllifolium. International Journal of Phytoremediation 13:229–244
Benizri E, Kidd PS (2018) The role of the rhizosphere and microbes associated with hyperaccumulator plants in metal accumulation. In: Van der Ent A, Echevarria G, Baker AJM, Morel JL (eds) Agromining: farming for metals: extracting unconventional resources using plants. Springer International Publishing, Cham
Berg G, Smalla K (2009) Plant species and soil type cooperatively shape the structure and function of microbial communities in the rhizosphere. FEMS Microbiol Ecol 68:1–13
Bordez L, Jourand P, Ducousso M, Carriconde F, Cavaloc Y, Santini S, Claverie JM, Wantiez L, Leveau A, Amir H (2016) Distribution patterns of microbial communities in ultramafic landscape: a metagenetic approach highlights the strong relationships between diversity and environmental traits. Mol Ecol 25:2258–2272
Brimecombe M, De Leij F, Lynch J (2001) The effect of root exudates on rhizosphere microbial populations. In: Pinton R, Varanini Z, Nannipieri P (eds) The rizosphere biochemistry and organic substances at the soil -plant Interface. Marcel Dekker, New York
Brooks RR (1987) Serpentine and its vegetation: a multidisciplinary approach. Dioscorides, Portland, OR
Brooks RR, Lee J, Reeves RD, Jaffre T (1977) Detection of nickeliferous rocks by analysis of herbarium specimens of indicator plants. J Geochem Explor 7:49–57
Brooks R, Shaw S, Marfil A (1981) Some observations on the ecology, metal uptake and nickel tolerance of Alyssum serpyllifolium subspecies from the Iberian Peninsula. Vegetatio 45:183–188
Cattani I, Capri E, Boccelli R, Del Re AAM (2009) Assessment of arsenic availability to roots in contaminated Tuscany soils by a diffusion gradient in thin films (DGT) method and uptake by Pteris vittata and Agrostis capillaris. Eur J Soil Sci 60:539–548
Costa R, Götz M, Mrotzek N, Lottmann J, Berg G, Smalla K (2006) Effects of site and plant species on rhizosphere community structure as revealed by molecular analysis of microbial guilds. FEMS Microbiol Ecol 56:236–249
Dakora FD, Phillips DA (2002) Root exudates as mediators of mineral acquisition in low-nutrient environments. Plant Soil 245:35–47
DeAngelis KM, Brodie EL, DeSantis TZ, Andersen GL, Lindow SE, Firestone MK (2008) Selective progressive response of soil microbial community to wild oat roots. ISME J 3:168–178
Delorme TA, Gagliardi JV, Angle JS, Chaney RL (2001) Influence of the zinc hyperaccumulator Thlaspi caerulescens J. & C. Presl. And the non metal accumulator Trifolium pratense L. on soil microbial populations. Can J Microbiol 47:773–776
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
Dias ACF, Hoogwout EF, MdC P e S, Salles JF, van Overbeek LS, van Elsas JD (2012) Potato cultivar type affects the structure of ammonia oxidizer communities in field soil under potato beyond the rhizosphere. Soil Biol Biochem 50:85–95
Essoussi I, Ghodhbane-Gtari F, Amairi H, Sghaier H, Jaouani A, Brusetti L, Daffonchio D, Boudabous A, Gtari M (2010) Esterase as an enzymatic signature of Geodermatophilaceae adaptability to Sahara desert stones and monuments. J Appl Microbiol 108:1723–1732
Fierer N, Bradford M, Jackson R (2007) Toward an ecological classification of soil bacteria. Ecology 88:1354–1364
Franco CMM, Labeda DP (2014) The order Pseudonocardiales. In: Rosenberg E, DeLong EF, Lory S, Stackebrandt E, Thompson F (eds) The prokaryotes. Springer, Berlin, Heidelberg, p 743–860
Galey ML, van der Ent A, Iqbal MCM, Rajakaruna N (2017) Ultramafic geoecology of south and Southeast Asia. Bot Stud 58:18
Garbeva P, van Veen JA, van Elsas JD (2004) Microbial diversity in soil: selection of microbial populations by plant and soil type and implications for disease suppressiveness. Annu Rev Phytopathol 42:243–270
Gomes NCM, Heuer H, Schonfeld J, Costa R, Mendonca-Hagler L, Smalla K (2001) Bacterial diversity of the rhizosphere of maize (Zea mays) grown in tropical soil studied by temperature gradient gel electrophoresis. Plant Soil 232:167–180
Gomes NCM, Kosheleva IA, Abraham WR, Smalla K (2005) Effects of the inoculant strain Pseudomonas putida KT2442 (pNF142) and of naphthalene contamination on the soil bacterial community. FEMS Microbiol Ecol 54:21–33
Gomes NCM, Landi L, Smalla K, Nannipieri P, Brookes PC, Renella G (2010) Effects of cd- and Zn-enriched sewage sludge on soil bacterial and fungal communities. Ecotoxicol Environ Saf 73:1255–1263
Gómez-Zotano J, Román-Requena F, Hidalgo-Triana N, Pérez-Latorre AV (2014) Biodiversity and conservation values of the serpentine ecosystems in Spain: Sierra Bermeja (Malaga Province). Boletin de la Asociacion de Geografos Espanoles 187–206+451–456
Goodfellow M (2014) The family Nocardiaceae. In: Rosenberg E, DeLong EF, Lory S, Stackebrandt E, Thompson F (eds) The prokaryotes: Actinobacteria. Springer, Berlin, Heidelberg, p 595–650
Gremion F, Chatzinotas A, Harms H (2003) Comparative 16S rDNA and 16S rRNA sequence analysis indicates that Actinobacteria might be a dominant part of the metabolically active bacteria in heavy metal-contaminated bulk and rhizosphere soil. Environ Microbiol 5:896–907
Gtari M, Essoussi I, Maaoui R, Sghaier H, Boujmil R, Gury J, Pujic P, Brusetti L, Chouaia B, Crotti E, Daffonchio D, Boudabous A, Normand P (2012) Contrasted resistance of stone-dwelling Geodermatophilaceae species to stresses known to give rise to reactive oxygen species. FEMS Microbiol Ecol 80:566–577
Haichar FZ, 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
Hartmann A, Schmid M, Dv T, Berg G (2009) Plant-driven selection of microbes. Plant Soil 321:235–257
Héry M, Nazaret S, Jaffré T, Normand P, Navarro E (2003) Adaptation to nickel spiking of bacterial communities in neocaledonian soils. Environ Microbiol 5:3–12
Hinsinger P, Courchesne F (2007) Biogeochemistry of metals and metalloids at the soil–root interface. Biophysico-chemical processes of heavy metals and metalloids in soil environments. John Wiley & Sons, Inc.
Idris R, Trifonova R, Puschenreiter M, Wenzel WW, Sessitsch A (2004) Bacterial communities associated with flowering plants of the Ni-hyperaccumulator Thlaspi goesingense. Appl Environ Microbiol 70:2667–2677
Kropf S, Heuer H, Grüning M, Smalla K (2004) Significance test for comparing complex microbial community fingerprints using pairwise similarity measures. J Microbiol Methods 57:187–195
Kuffner M, Puschenreiter M, Wieshammer G, Gorfer M, Sessitsch A (2008) Rhizosphere bacteria affect growth and metal uptake of heavy metal accumulating willows. Plant Soil 304:35–44
Kušar D, Avguštin G (2012) Optimization of the DGGE band identification method. Folia Microbiol 57:301–306
Lipman CB (1926) The bacterial flora of serpentine soils. J Bacteriol 12:315–318
López González G (1975) Contribución al estudio florístico y fitosociológico de Sierra de Aguas. Acta Botanica Malacitana 1:81–205
Lopez S, Piutti S, Vallance J, Morel J-L, Echevarria G, Benizri E (2017) Nickel drives bacterial community diversity in the rhizosphere of the hyperaccumulator Alyssum murale. Soil Biol Biochem 114:121–130
Ma Y, Oliveira RS, Freitas H, Zhang C (2016) Biochemical and molecular mechanisms of plant-microbe-metal interactions: relevance for phytoremediation. Front Plant Sci 7:918
Marschner P, Crowley D, Yang CH (2004) Development of specific rhizosphere bacterial communities in relation to plant species, nutrition and soil type. Plant Soil 261:199–208
McCaig AE, Glover LA, Prosser JI (1999) Molecular analysis of bacterial community structure and diversity in unimproved and improved upland grass pastures. Appl Environ Microbiol 65:1721–1730
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
Menezes de Sequeira E (1969) Toxicity and movement of heavy metals in serpentinic soils (North-Eastern Portugal). Agron Lusit 30:115–154
Menezes de Sequeira E, Pinto da Silva AR (1992) Ecology of serpentinized areas of north-east Portugal. In: Roberts BA, Proctor J (eds) The ecology of areas with serpentinized rocks. Springer, Dordrecht, p 169–197
Mengoni A, Barzanti R, Gonnelli C, Gabbrielli R, Bazzicalupo M (2001) Characterization of nickel-resistant bacteria isolated from serpentine soil. Environ Microbiol 3:691–698
Mengoni A, Grassi E, Barzanti R, Biondi EG, Gonnelli C, Kim CK, Bazzicalupo M (2004) Genetic diversity of bacterial communities of serpentine soil and of rhizosphere of the nickel-hyperaccumulator plant Alyssum bertolonii. Microb Ecol 48:209–217
Mengoni A, Schaat H, Vangronsveld J (2010) Plants as extreme environments? Ni-resistant bacteria and Ni-hyperaccumulators of serpentine flora. Plant Soil 331:5–16
Muehe EM, Weigold P, Adaktylou IJ, Planer-Friedrich B, Kraemer U, Kappler A, Behrens S (2015) Rhizosphere microbial community composition affects cadmium and zinc uptake by the metal-hyperaccumulating plant Arabidopsis halleri. Appl Environ Microbiol 81:2173–2181
Normand P, Daffonchio D, Gtari M (2014) The family Geodermatophilaceae. In: Rosenberg E, DeLong EF, Lory S, Stackebrandt E, Thompson F (eds) The prokaryotes: Actinobacteria. Springer, Berlin, Heidelberg, p 361–379
Oline D (2006) Phylogenetic comparisons of bacterial communities from serpentine and non serpentine soils. Appl Environ Microbiol 72:6965–6971
Oren A, Xu X-W (2014) The family Hyphomicrobiaceae. In: Rosenberg E, DeLong EF, Lory S, Stackebrandt E, Thompson F (eds) The prokaryotes: Alphaproteobacteria and Betaproteobacteria. Springer, Berlin Heidelberg, Berlin, Heidelberg
Pal A, Dutta S, Mukherjee PK, Paul AK (2005) Occurrence of heavy metal-resistance in microflora from serpentine soil of Andaman. J Basic Microbiol 45:207–218
Philippot L, Raaijmakers JM, Lemanceau P, Van Der Putten WH (2013) Going back to the roots: the microbial ecology of the rhizosphere. Nat Rev Microbiol 11:789–799
Pollard AJ, Reeves RD, Baker AJM (2014) Facultative hyperaccumulation of heavy metals and metalloids. Plant Sci 217–218:8–17
Proctor J (1999) Toxins, nutrient shortages and droughts: the serpentine challenge. Trends Ecol Evol 14:334–335
Rivas R, Velázquez E, Willems A, Vizcaíno N, Subba-Rao NS, Mateos PF, Gillis M, Dazzo FB, Martínez-Molina E (2002) A new species of Devosia that forms a unique nitrogen-fixing root-nodule symbiosis with the aquatic legume Neptunia natans (L.f.) Druce. Appl Environ Microbiol 68:5217–5222
Rivas R, Willems A, Subba-Rao NS, Mateos PF, Dazzo FB, Kroppenstedt RM, Martínez-Molina E, Gillis M, Velázquez E (2003) Description of Devosia neptuniae sp. nov. that nodulates and fixes nitrogen in symbiosis with Neptunia natans, an aquatic legume from India. Syst Appl Microbiol 26:47–53
Sandaa RA, Torsvik V, Enger Ø, Daae FL, Castberg T, Hahn D (1999) Analysis of bacterial communities in heavy metal-contaminated soils at different levels of resolution. FEMS Microbiol Ecol 30:237–251
Sandaa RA, Torsvik V, Enger Ø (2001) Influence of long-term heavy-metal contamination on microbial communities in soil. Soil Biol Biochem 33:287–295
Schmalenberger A, Tebbe CC (2003) Bacterial diversity in maize rhizospheres: conclusions on the use of genetic profiles based on PCR-amplified partial small subunit rRNA genes in ecological studies. Mol Ecol 12:251–262
Schwartz C, Morel JL, Saumier S, Whiting SN, Baker AJM (1999) Root development of the zinc-hyperaccumulator plant Thlaspi caerulescens as affected by metal origin, content and localization in soil. Plant Soil 208:103–115
Segura A, Rodríguez-Conde S, Ramos C, Ramos JL (2009) Bacterial responses and interactions with plants during rhizoremediation. Microb Biotechnol 2:452–464
Smalla K, Wieland G, Buchner A, Zock A, Parzy J, Kaiser S, Roskot N, Heuer H, Berg G (2001) Bulk and rhizosphere soil bacterial communities studied by denaturing gradient gel electrophoresis: plant-dependent enrichment and seasonal shifts revealed. Appl Environ Microbiol 67:4742–4751
Steer J, Harris JA (2000) Shifts in the microbial community in rhizosphere and non-rhizosphere soils during the growth of Agrostis stolonifera. Soil Biol Biochem 32:869–878
Thijs S, Langill T, Vangronsveld J (2017) The bacterial and fungal microbiota of hyperaccumulator plants: small organisms, large influence. Adv Bot Res 83:43–86
Touceda-González M, Prieto-Fernández Á, Renella G, Giagnoni L, Sessitsch A, Brader G, Kumpiene J, Dimitriou I, Eriksson J, Friesl-Hanl W, Galazka R, Janssen J, Mench M, Müller I, Neu S, Puschenreiter M, Siebielec G, Vangronsveld J, Kidd PS (2017) Microbial community structure and activity in trace element-contaminated soils phytomanaged by gentle remediation options (GRO). Environ Pollut 231:237–251
van der Ent A, Baker AJM, van Balgooy MMJ, Tjoa A (2013) Ultramafic nickel laterites in Indonesia (Sulawesi, Halmahera): mining, nickel hyperaccumulators and opportunities for phytomining. J Geochem Explor 128:72–79
Visioli G, D'Egidio S, Sanangelantoni AM (2015) The bacterial rhizobiome of hyperaccumulators: future perspectives based on omics analysis and advanced microscopy. Front Plant Sci 5:752
Wenzel WW, Bunkowski M, Puschenreiter M, Horak O (2003) Rhizosphere characteristics of indigenously growing nickel hyperaccumulator and excluder plants on serpentine soil. Environ Pollut 123:131–138
Whittaker RH (1954) The ecology of serpentine soils. Ecology 35:258–288
Acknowledgements
This research was funded by the Ministerio de Economía, Industria y Competitividad and FEDER (FACCE Surplus project Agronickel (ID71) PCIN-2017-028 and CTM2015-66439-R).
Author information
Authors and Affiliations
Corresponding author
Additional information
Responsible Editor: Antony Van der Ent
Electronic supplementary material
ESM 1
(DOCX 1454 kb)
Rights and permissions
About this article
Cite this article
Touceda-González, M., Kidd, P.S., Smalla, K. et al. Bacterial communities in the rhizosphere of different populations of the Ni-hyperaccumulator Alyssum serpyllifolium and the metal-excluder Dactylis glomerata growing in ultramafic soils. Plant Soil 431, 317–332 (2018). https://doi.org/10.1007/s11104-018-3767-6
Received:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1007/s11104-018-3767-6