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Responses of microbial communities to drought stress and humic acid fertilizer in maize

  • # Authors contributed equally: Mingfei Sun, Lin Zhang

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  • Humic acid (HA) fertilizer was previously considered to be able to enlarge the effective absorption area of roots and promote plant root growth. Recent evidence suggests that certain root-associated microbes might be able to mitigate the negative responses of drought stress. In this study, the abundance and diversity of root-associated bacterial communities under HA fertilizer and drought stress treatments were explored. Rhizosphere microbiomes of three groups (HA, drought, and control) and microbiomes from bulk soil were collected during the flowering period in maize. Using Illumina Hiseq2500, the bacterial data for the V4 region of the 16S rRNA gene was obtained. Analysis of the sequencing data from 40 bacteria phyla revealed the abundance levels of the 12 bacterial communities were significantly different in the rhizosphere of the drought-treated samples compared to the controls. Two of these phyla, Actinobacteria and Bacteroidetes, were also significantly enriched or depleted in HA fertilizer-treated samples. The results underscore the dynamics of the rhizosphere microbiomes under drought stress and indicate the association between HA and rhizosphere microbiomes. However, more studies are warranted to understand the effect of HA fertilizer and beneficial microbiomes in responding to drought.
  • The contamination of ecosystems with heavy metals has become a concern due to their toxicity to biota and also due to them being permanently immobilised in soil components. Moreover, metals influence soil microbes by decreasing their population size, diversity and biochemical activity, thereby altering the structure of soil microbial communities[13]. Exposure to metals can also result in the establishment of tolerant microbial populations in soil, with important roles in the ecosystem.

    Arsenic (As) is a ubiquitous trace metalloid present in almost all environments and widely distributed in soil and water[4]. The amounts of As in non-polluted soils are usually lower than 10 mg·kg−1[5]. The occurrence of high levels of As in soils results from human activities, such as mining and smelting processes, in addition to the use of As-based compounds such as insecticides, herbicides, fungicides, algaecides, sheep dips, wood preservatives, dyestuffs and feed additives[5]. Natural sources of As comprise volcanic activities, wind-born soil particles, sea salt sprays and microbial volatilization[6,7].

    Currently available tools for the remediation of soils polluted with metals or metalloids are costly, time consuming, can be dangerous to people and result in the production of secondary waste[8,9]. Phytoremediation has become known as a promising eco-remediation technology by which plants and their associated microbiota are used to eradicate contaminants from polluted soils[10,11], acting through phytoextraction, stabilization, immobilization, volatilization and rhizofiltration[1215].

    Soils contaminated by heavy metals frequently have low percentages of organic matter and available nitrogen, limiting plant development[16]. Legume plants and their associated root nodule bacteria (rhizobia) are essential components of the biogeochemical cycles in both natural and agricultural ecosystems. We consider that this association could aid rehabilitation of disturbed areas by adding fixed atmospheric nitrogen that can be used by other plants (non-legumes). However, the growth and development of many plant species, including legumes, is affected by the occurrence of high amounts of toxic chemical elements such as As.

    Compared to other legumes, Lotus species have a higher potential for adaptability to abiotic stresses, frequently surviving in adverse conditions such as those found in polluted soils[17,18]. Lotus is a cosmopolitan genus with two main centers of diversity, the Mediterranean region (including portions of Europe, Africa, and Western Asia) and Western North America[19,20]. The genus Lotus comprises about 100 annual and perennial species[18]. However, only a small number of Lotus species, mainly of agronomic interest, have been studied in relation to their symbionts[21,22]. The species of Lotus that have been domesticated and improved by selection and plant breeding are: L. corniculatus, L.; L. uliginosus Schkuhr., also denoted as L. pedunculatus; L. glaber Mill., also denoted as L. tenuis; L. subbiflorus Lagasca and L. ornithopodioides[18]. On the other hand, L. japonicus is used as a model for genetic and molecular studies[2325].

    L. uliginosus is a perennial legume also used as a tropical forage with a high productive capacity and is recommended for extensive livestock areas in countries of the South American cone and especially in Chile, Brazil and Uruguay. This species was also naturalised in Argentina. Its use is in cattle and sheep rearing and fattening processes and also for soil remediation. It thrives in varied conditions, especially low fertility and moisture as well as soil acidity. It is a species with high spring-summer-autumn forage potential comparable to other traditionally used legumes. This specie has also a outstanding nutritional value, with the presence of condensed tannins, giving it additional nutritional advantages[26]. Due to its importance several authors have studied the diversity and phylogenetic relationships of root nodule bacteria of L. uliginosus collected from fields in different countries, such as Uruguay[27], Portugal[28] and Belgium[29]. These authors indicated that those strains were affiliated with Bradyrhizobium japonicum or Bradyrhizobium sp.

    The goal of this research was to investigate a symbiotic legume system effective in nitrogen fixation in a polluted site for possible use during remediation of such soils. In view of the future use of legumes for soil improvement, it is essential to assess the effect of As on the functioning of the symbiosis. Therefore, the objective of this work was also to study the rhizobial population associated with L. uliginosus growing in As contaminated soils with regards to genetic diversity, efficiency of nitrogen-fixation and tolerance to As (Supplemental Fig. S1).

    Bacteria were isolated from root nodules of Lotus uliginosus growing under field conditions on an industrially contaminated soil, in the centre of Portugal (GPS: 40.778978329572496, −8.59899224806268), containing high levels of arsenic (1.5 × 103 mg·kg−1). Six plants were collected and 5−6 nodules were randomly taken from each plant. The nodules were carefully detached from the roots, surface sterilized (0.25% solution of HgCl2)[28] and washed extensively with sterile water. Afterwards, nodules were individually crushed and spread on a plate with yeast-mannitol agar (YMA) supplemented with congo red dye[30]. Plates were incubated at 28 ºC in the dark for 7-10 d. The isolate purity was checked by examining colony morphology and congo red absorption. Isolates were also cultivated in yeast-mannitol agar (YMA) supplemented with bromothymol blue (BTB)[30]. Isolates were kept at 4 ºC.

    ERIC (Enterobacterial Repetitive Intergenic Consensus) - PCR[31] was used to assess the genetic diversity of 22 isolates obtained from nodules of L. uliginosus as described above. This procedure is used to distinguish strains that are taxonomically very close and has been recognized as appropriate for assessing rhizobial diversity[32,33]. DNA was extracted from bacterial liquid cultures using the Aqua pure genomic DNA extraction kit from Bio-Rad, following the kit protocol specifications. ERIC fingerprints were generated using the primers ERIC1R and ERIC2 previously reported by de Bruijn and Versalovic et al.[31,34]. Matrices of the Dice coefficient were calculated and cluster analysis was performed using the UPGMA (Unweighted Pair Group with Arithmetic Average) algorithm and the program Free Tree[35,36]. The program Tree View (PHILIP) was used for the construction of dendrograms and evaluation of the respective genetic relationships.

    The 16S rRNA region was amplified in seven selected isolates using 41F and 1488R primers as described by Weisburg et al.[37]. PCR reactions primers were used at a concentration of 5 μM, together with 15.8 μl of Qiagen kit Taq mix solution (2.5 U of Taq polymerase, 1.5 mM of MgCl2, 200 μM of the different dNTP), and approximately 20 ng of genomic DNA, in a final volume of 20 μl. Amplifications were performed following the conditions of Weisburg et al.[37], using an Eppendorf Mastercycler Gradient thermocycler. PCR products of the amplified 16S rRNA region, with an expected size of about 1500 bp, were confirmed by agarose gel electrophoresis. These products were sequenced, with the same primers used for PCR amplification, with an ABI 3730 XL automated sequencer, by STAB VIDA, Caparica, Portugal. Obtained nucleotide sequences were subjected to quality control and edited as necessary using the DNA chromatogram files in Chromas Lite program (version 2.1.1). Homologous sequences were searched in NCBI (National Center for Biotechnology Information) GenBank database (www.ncbi.nlm.nih.gov) using BLASTn tool[38]. Nucleotide sequences were aligned using ClustalW within the MEGA 7 platform. A neighbor-joining phylogenetic tree was constructed using the p-distance model and tested with 1000 bootstrap replication within the MEGA 7 platform[39].

    Sequences obtained previously were deposited in the NCBI database. Below is a list of all accession codes presented next to the correspondent strain:

    Isolate 8: OQ145681

    Isolate 10: OQ145686

    Isolate 12: OQ145682

    Isolate 15: OQ145683

    Isolate 21: OQ145684

    Isolate 23: OQ145685

    Isolate 24: OQ145687

    Arsenic tolerance of isolates was assessed by evaluating growth inhibition in the presence of different levels of As. For each isolate, a cell suspension was prepared in sterile water and 30 µL of a pre-washed re-suspended aliquot (106 cells mL−1) was inoculated in 5 mL of yeast-mannitol (YM) liquid medium containing increasing concentrations of arsenic(III) chloride (AsCl3) (5, 10, 50 and 200 mg of As mL−1). Tubes were kept at 28 ºC on an orbital shaker at 100-rev min−1 for 72 h. Isolate growth was evaluated by a once per day measurement of the optical density at 600 nm. Two replicas for each strain and each concentration were carried out. Finally, the growth of the bacterial isolates was classified into three groups according to the percentage of growth inhibition: sensitive (100%−80%), moderately tolerant (80%−60%) and tolerant (< 60%). Cell suspensions prepared without the addition of As were also included as controls for each isolate.

    Lotus uliginosus cv. Sunrise seeds were surface sterilized and rinsed extensively with sterile distilled water according to[30]. Next, seeds were hydrated for 1−2 h in sterilized water and moved to 0.8% w v−1 agar-water plates for 1−2 days until germination. The pre-germinated seeds were moved to slants containing 50 mL of N-free Jensen plant medium[40]. Seedlings were allowed to establish in this nutrient medium before addition of bacterial inocula and As treatments. Each isolate was inoculated by applying 1 mL of bacterial suspension (approximately 108 bacterial cells in 1/4 Jensen medium) on the roots of each seedling (3 replicates per isolate). Arsenic was added to liquid Jensen medium (1/4 diluted) as AsCl3, in order to produce concentrations of 0.5, 5, 10, 20, 100 and 200 mg·mL−1 of As. Plants were supplemented with each As concentration (three replicates per each treatment). Furthermore, controls only with nitrogen (TN) and without nitrogen and not inoculated (T0) were prepared as described by Soares et al. [22]. Plants were incubated for six weeks in a controlled-environment chamber with 16 h light/8 h dark cycle at 23 ºC (day)/18 ºC (night). Plants were observed for nodulation after three and six weeks, and were harvested after six weeks of growth. The colour of the interior of the nodules was also observed and correlated with the presence/absence of leghemoglobin depending on whether they were pink or not, i.e., root nodules coloured pink by leghaemoglobin are caused by a nitrogen-fixing symbiotic relationship between the plant and beneficial bacteria (rhizobia)[41].

    Finally, the shoots were dried at 80 ºC for two days. The dry weight (aerial biomass) of inoculated plants (X) was used to determine the index of effectiveness (Es) as described by Ferreira & Marques[42]. TN and T0 represent the dry weight of plants from nitrogen control and from non-inoculated plants, respectively: =XT0TNT0×100% .

    Data were analysed by one-way analysis of variance (ANOVA) with software STATISTICA version 10 (StatSoft), using the Tukey’s honestly significance difference (HSD) test at P ≤ 0.05.

    In order to address the objective of this research, we isolated bacterial strains from L. uliginosus growing under field conditions in an arsenic contaminated site in the central region of Portugal. In this context, 22 isolates were obtained from root nodules of L. uliginosus. All isolates were slow growers, showed little congo red absorption, and had an alkaline reaction on BTB, indicated by a blue colour, which is usually produced by Bradyrhizobium spp.[30] (Supplemental Fig. S2). Congo red is often incorporated in culture media for isolating rhizobia or for testing the purity of rhizobia cultures. Rhizobia typically do not absorb congo red or absorb it weakly, while other bacteria absorb it strongly.

    The assessment of genotypic diversity of the natural population nodulating L. uliginosus was achieved by ERIC-PCR. The analysis of the fingerprinting patterns of each isolate showed the existence of several clusters (Fig. 1). This dendrogram was used to determine the similarities among isolates. Results showed the presence of a high genetic diversity in the population, despite the high contamination by As in the original soil. These isolates showed multiple fingerprinting patterns and no single dominant genotype was apparent from our results.

    Figure 1.  Dendrogram showing the diversity of root nodule bacteria (Bradyrhizobium sp.) isolated from L. uliginosus, derived from ERIC-PCR fingerprints using UPGMA method, at 85% similarity.

    Results obtained for As tolerance after 72 h (Fig. 2, Supplemental Fig. S3 and Table 1) showed that, for the highest concentration tested (200 mg of As mL−1), most isolates were considered As sensitive (percentage of growth inhibition (GI) of 80%−100%). Only two isolates, 13 and 12, were considered as moderately tolerant (GI 60%−80%) and tolerant (GI < 60%), respectively. On the other hand, for 50 mg·mL−1 of As about 36% of isolates (isolates 8, 12, 13, 14, 15, 16, 17 and 23) were tolerant, and 18% were considered moderately tolerant (isolates 7, 18, 19 and 22). However, an inverse situation was verified for 10 and 5 mg·mL−1 of As, where no isolates were found sensitive. While for 10 mg·mL−1 64% of the isolates were tolerant, for the lowest concentration of As used, 5 mg·mL−1, a large majority of isolates (95%) was considered as tolerant.

    Figure 2.  Tolerance of the L. uliginosus isolates to different As concentrations (5, 10, 50 and 200 mg·mL−1). Isolates were classified according to the percentage of growth inhibition relative to controls grown in the absence of As, being considered sensitive (80%−100%), moderately tolerant (60%−80%) and tolerant (< 60%). Stacked-columns indicate the percentage of isolates in the three tolerance classes for each As concentration.
    Table 1.  Nodulation phenotype of L. uliginosus plants inoculated with the different isolates upon different As concentrations.
    IsolatesAs concentration (mg·mL−1)
    00.551020
    3 w6 w3 w6 w3 w6 w3 w6 w3 w6 w
    8++++++++
    ++++++++
    ++++++++
    10++±±
    ++±±
    ++±±
    12++++++±±
    ++++++±±
    ++++++±±
    15+++++++
    +++++++++
    +++++++++
    21+++++++
    +++++++
    +++++++
    23++++++
    +++++++
    ++++++++
    24++++++
    ++++++
    ++++
    w, incubation weeks; +, presence of nodules; −, absence of nodules; ±, presence of small and white nodules.
     | Show Table
    DownLoad: CSV

    For the evaluation of the effects of As on the symbiosis with L. uliginosus, seven isolates (namely 8, 10, 12, 15, 21, 23 and 24) belonging to different clusters (Fig. 1) and with different levels of tolerance to As (Supplemental Fig. S3) were chosen to inoculate L. uliginosus cv. Sunrise seedlings. Assays were performed with six As concentrations (0.5, 5, 10, 20, 100 and 200 mg of As mL−1) and results were recorded three and six weeks after the addition of bacterial inocula and As to the seedlings. Briefly, plants did not tolerate the highest As concentrations used, 100 and 200 mg·mL−1, and died one week after As addition. Arsenic toxicity affected the symbiosis and the different isolates also showed different symbiotic performances as the arsenic concentration increased up to 20 mg·mL−1 (Fig. 3).

    Figure 3.  Effect of As on the symbiosis. Values represent the average of the index of effectiveness (Es) ± SD of L. uliginosus plants inoculated with the bacterial isolates and grown with different As concentrations (0, 0.5, 5, 10 and 20 mg·mL−1). Different letters express significant differences between plants inoculated with each isolate at several As concentrations according to Tukey's HSD test at at P < 0.05.

    One of the effects of As in the symbiosis was demonstrated by the delay in nodulation (Table 1), especially for higher concentrations of As tested, when compared to the controls without the addition of As. A more drastic effect was the impairment of the symbiosis by the lack of nodulation. For the highest As concentration allowing plant survival, 20 mg·mL−1, only plants inoculated with isolates 15, 21 and 23 presented pink root nodules (i.e., functional) at least in one of the replicates. In this case, nodulation was delayed, but the symbiosis remained effective in some replicates, depending on the isolate tested. Plants inoculated with isolates 8 and 12 did not show delayed nodulation but nodules were only formed at As concentrations up to 10 mg·mL−1. However, at this concentration plants inoculated with isolate 12 had small and white nodules (i.e., ineffective). Nodulation was also observed (with the presence of pink nodules) in plants inoculated with isolate 24 until 10 mg of As mL−1 but with a delay of 3 weeks. Plants inoculated with isolate 10 and growing with 0.5 and 5 mg of As mL−1 showed also a delay in nodulation and nodules were small and white. This nodulation phenotype indicates an ineffective symbiosis, inversely to what happened with plants grown without the addition of As which had pink nodules.

    These plant-inoculation experiments using several As concentrations were also performed to evaluate the symbiotic effectiveness, by determining the shoot dry weight of L. uliginosus plants after 6 weeks of growth. The As concentrations used in these experiments were higher than those usually present in groundwater used for irrigation and in soils in various countries, e.g. 3,10 µg·L−1 and 22 mg·kg−1, respectively, in Argentina, or 3,700 µg·L−1 and 196 mg·kg−1, respectively, in India[43]. All the isolates chosen for these experiments were considered effective in the absence of As and the respective indices of effectiveness (Es), under these conditions, ranged between 35% and 55% (Fig. 3). These isolates also showed different levels of symbiotic performance and indices of effectiveness upon different As concentrations. In general, results were congruent with the observed nodulation phenotypes (Table 1). Isolates 15, 21 and 23 were able to establish an efficient symbiosis with the host plant at 20 mg of As mL−1. Plants inoculated with isolate 15 showed an increase in the aerial part and the consequent increase in the indices of effectiveness at the concentration of 0.5 mg As mL−1, which was near 75% and the highest Es of these experiments. This strain was therefore considered highly effective in the presence of As (0.5 mg·mL−1). However, for the other two isolates, 21 and 23 respectively, no significant differences were found between the various treatments, including the control without As addition, and their respective indices of effectiveness were lower but still considered as effective in nitrogen fixation. On the other hand, isolates 8, and 24, were able to establish an efficient symbiosis until the concentration of 10 mg of As mL−1. For the first isolate, no significant differences were found between the various treatments (0.5, 5 and 10 mg As mL−1) including the control without addition of As. The indices of effectiveness of plants inoculated with isolate 12 and with 0.5 and 5 mg of As mL−1 did not show significant differences between each other and the control (without As addition). However, for 10 mg·mL−1 of As the index of effectiveness was very low (19%) and the symbiosis was considered as ineffective, the plants presenting small and white nodules as previously mentioned. Lastly, plants inoculated with isolate 10 had a very weak performance in the presence of As (with plants having small and white nodules) and were only able to establish an efficient symbiosis when As was completely absent, showing indices of effectiveness significantly different from all the remaining treatments.

    Interestingly, among the isolates that were able to establish an efficient symbiosis while sustaining the highest As concentration tolerated by plants, 20 mg·mL−1 (isolates 15, 21 and 23), isolate 15 was also among those with higher indices of effectiveness in the absence of As. Moreover, this isolate had the highest Es shown, near 75% at 0.5 mg·mL−1 of As being considered as highly effective in nitrogen fixation.

    Aligned sequences of the partial 16S rRNA region were used to construct the phylogenetic tree shown in Fig. 4. The isolates from this study were all clustered with Bradyrhizobium spp. Sequences obtained from isolates 8, 10, 15, 23 and 24 shared 100% sequence identity and the closest strains were Bradyrhizobium spp. isolated from Lotus uliginosus (Bradyrhizobium sp. 3LBC, 8LBI, SEMIA 839 and NZP2309[21,28]), Cytisus triflorus (Bradyrhizobium sp. CTS12), Cytisus scoparius (Bradyrhizobium genosp. AD Cs6020[44]) Vigna unguiculata L. (Bradyrhizobium sp. VUPMI37) and Ulex europaeus (Bradyrhizobium sp. ICMP 12674[45]). Isolates 12 and 21 shared 100% sequence identity and the closest strains were Bradyrhizobium spp. isolated from Glycine max (Bradyrhizobium japonicum GI-4 and J5 and Bradyrhizobium sp. 323S2[46,47]), Pigeonpea (Bradyrhizobium sp. RP6) and Erythrina brucei (Bradyrhizobium shewense ERR11T [48]).

    Figure 4.  Phylogeny of the partial 16S rRNA gene with a total of 1189 aligned positions. Confidence bootstrap values are presented near each node. NCBI GenBank accession codes are presented next to each strain. Isolates obtained in this work are in bold. Rhizobium leguminosarum USDA 2370T (U29386) was selected as an outgroup.

    Biological nitrogen fixation, including the contribution made by symbioses in the root nodules of legumes, supplies a large proportion of the nitrogen that increases soil fertility in natural and agro-ecosystems[49]. In recent years, the selection of symbiotic or free-living plant growth promoting rhizobacteria with remediation abilities has gained prominence, since such strains could help plants to grow in polluted soils or could even limit the inclusion of contaminants into plant tissues[50]. In particular, the rhizobium-legume symbiotic interaction has been highlighted as a promising tool to be used for bioremediation of As and heavy metals in soils[5155].

    It has been reported by several authors that associations between plants and microorganisms increase the bioremediation potential of plants[5658]. In this study we found a large diversity of root nodule bacteria isolated from L. uliginosus growing in soils contaminated with As in Portugal. These results are different from previous data indicating a lack of genetic diversity in the population of Rhizobium isolated from Trifolium sp. in an analogous area contaminated by heavy metals[59,60]. However, similar results have been reported by Carrasco et al.[51] and also by Delorme et al.[61], who observed that the presence of rhizobial genotypes in soils with high heavy metal concentration were not different from those existing in soils with low metal levels. These authors showed that the rhizobial population was very diverse and the presence of heavy metals did not lead to a decrease in diversity. According to Rangel et al.[62], a wide diversity of bacteria resistant to As was verified, including several rhizobial genera such as Azorhizobium, Mesorhizobium, Rhizobium, Burkholderia (now either in Paraburkholderia or Trinickia)[63]. Therefore, biological nitrogen fixation has a great potential to be used in the future for phytostabilization purposes, given the high number of host legume species of the mentioned rhizobia that can be tested in the field and also because legumes accumulate heavy metals mainly in roots and show a low level of metal translocation to the shoot[55].

    In this study, we have isolated bacteria from root nodules of L. uliginosus growing in soil contaminated by arsenic. These bacteria were morphologically characterized, by growth rate and reaction on BTB, as identical to those strains belonging to the genus Bradyrhizobium. Moreover, L. uliginosus root nodule bacteria seem to be constrained to B. japonicum and Bradyrhizobium sp. symbionts, rarely harbouring endophytic bacteria[27,28]. These results were confirmed for a set of seven isolates (namely, 8, 10, 12, 15, 21, 23 and 24) used to test the effects of As in the symbiosis, which were molecularly identified as Bradyrhizobium spp., using 16S rRNA region sequencing. A detailed phylogenetic analysis comprising appropriate molecular markers[22] could reveal if these isolates represent novel species or described ones, and compare them with the ones already described taxonomically from L. uliginosus that did not originate from As-contaminated soils[18, 24].

    In the tests performed to assess As tolerance, it was found that all the isolates were able to grow at concentrations of 5 and 10 mg As mL−1 revealing higher tolerance than observed by Deepika et al.[64] for Rhizobium radiobacter (strain VBCK1062) when using 10 mM As in YM liquid medium.

    In the tests carried out with L. uliginosus plants, the effects of As on nodulation and growth were recorded three and six weeks after seeding. The number of nodules and growth of the plants were affected by the presence of As in the medium, producing a severe effect in the early stage of nodulation. This could be due to the effect of As on rhizobia survival and initial tolerance[65]. The negative effects of As in the nodulation process found in our study are similar to those described by several authors. In fact, Reichman[66] observed that treatment with As induced a significant decrease in the total and effective number of nodules in soybean. Also, Pajuelo et al.[67] found important proof of damage by As on roots as well as a reduction of root hair number, which was associated with a decrease of around 90% in rhizobial infection number in plants of Medicago sativa. Moreover, the reduction in nodulation observed in legumes exposed to other metals, such as Cu2+, Cd2+ and Hg2+, was also attributed to the damage of metals on root hairs[68]. On the other hand, Bustingorri et al.[69] demonstrated that As causes membrane damage and decreases the chlorophyll content. This metalloid has no known biological function and fundamentally disrupts the cell growth and metabolism of living cells[64]. The adaptive tolerance response occurring in some rhizobial strains to the perturbation caused by As contamination, was also reported by others[47]. Our results confirmed that some Bradyrhizobium isolates tested for the evaluation of the effects of As in the symbiosis showed capacity to establish an efficient symbiosis, even in the presence of the highest As concentration (20 mg·mL−1), mainly isolate 15. This isolate had high effectiveness when compared to the As-free inoculated controls and to the TN and T0 controls.

    Biological nitrogen fixation, including the contribution made by symbioses with legumes, supplies a large proportion of the nitrogen that increases soil fertility in natural and agro-ecosystems[49]. In recent years, the selection of symbiotic or free-living plant growth promoting rhizobacteria with remediation abilities has gained prominence, since such strains could help plants to grow in polluted soils or could even limit the inclusion of contaminants into plant tissues[50]. In particular, the Rhizobium-legume symbiotic interaction has been highlighted as a promising tool to be used for bioremediation of As and heavy metals in soils[5154]. In this study, we have isolated bacteria from L. uliginosus with the capacity to establish an efficient symbiosis, even in the presence of high concentrations of As.

    We consider Bradyrhizobium isolate 15, as the most promising to be tested in situ for their applicability for phytoremediation in sites polluted by this metalloid.

    The main impact of this work is the possible use of autochthonous legume plants and their micro-symbionts, such as the symbioses with L. uliginosus and Bradyrhizobium, for the phytostabilization of contaminated soils, helping its fertilization. Results reveal that root nodule bacteria isolated from L. uliginosus growing in polluted soils, mainly those tolerant to the highest concentration of arsenic, can be symbiotically effective upon high As concentrations. This can be a very interesting aspect and can be used in the future, in bioremediation experiments on contaminated soils using native legumes, since it could have a positive ecological impact in those sites. We consider that the dual function of As bioremediation plus soil nitrogen enhancement can be achieved by effective symbiotic nodulation in affected As-metal-soils. Such a tolerant and functional symbiosis can support vegetation cover, stabilizing As-contaminated soils, and consequently it could be a smart practice for phytostabilization.

    This work was supported by the cooperation project between Portugal and Argentina: 'The genus Lotus and their utilization for the restoration of soils contaminated with heavy metals. The biochemistry and their symbionts', FCT/DREBM 00264, Proc. 4.1.3 and also by the European project: 'Raising the bio-based industrial feedstock capacity of marginal Lands (Margin Up)', nº101082089.

  • The authors declare that they have no conflict of interest.

  • Supplementary Fig. S1 Heatmap of the bacteria phylum (A) as well as Shannon’s diversity (B&C) represented in the control soil (S), control (C), drought (D) and HA (H) rhizosphere microbiome.
    Supplementary Fig. S2 Wilcoxon Signed Rank Test for percent relative abundance of the abundant bacterial phylum in Control Soil, Control rhizosphere (Control Rhizo) and drought rhizosphere (Drought Rhizo) groups.
    Supplementary Fig. S3 Summary statistics of phenotypes.
    Supplementary Fig. S4 Wilcoxon Signed Rank Test for relative abundance of the abundant bacterial phyla in control soil, control rhizosphere and Humic Acid rhizosphere groups.
    Supplementary Fig. S5 Summary statistics of relative abundance of the top 12 of the most abundant baterial families.
    Supplementary Fig. S6 Summary analyses of LEfSe among control soil (S), control (C), drought (D) and HA (H) rhizosphere conditions.
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  • Cite this article

    Sun M, Zhang L, Zhu H, Kang C, Li X, et al. 2024. Responses of microbial communities to drought stress and humic acid fertilizer in maize. Genomics Communications 1: e006 doi: 10.48130/gcomm-0024-0006
    Sun M, Zhang L, Zhu H, Kang C, Li X, et al. 2024. Responses of microbial communities to drought stress and humic acid fertilizer in maize. Genomics Communications 1: e006 doi: 10.48130/gcomm-0024-0006

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Responses of microbial communities to drought stress and humic acid fertilizer in maize

Genomics Communications  1 Article number: e006  (2024)  |  Cite this article

Abstract: Humic acid (HA) fertilizer was previously considered to be able to enlarge the effective absorption area of roots and promote plant root growth. Recent evidence suggests that certain root-associated microbes might be able to mitigate the negative responses of drought stress. In this study, the abundance and diversity of root-associated bacterial communities under HA fertilizer and drought stress treatments were explored. Rhizosphere microbiomes of three groups (HA, drought, and control) and microbiomes from bulk soil were collected during the flowering period in maize. Using Illumina Hiseq2500, the bacterial data for the V4 region of the 16S rRNA gene was obtained. Analysis of the sequencing data from 40 bacteria phyla revealed the abundance levels of the 12 bacterial communities were significantly different in the rhizosphere of the drought-treated samples compared to the controls. Two of these phyla, Actinobacteria and Bacteroidetes, were also significantly enriched or depleted in HA fertilizer-treated samples. The results underscore the dynamics of the rhizosphere microbiomes under drought stress and indicate the association between HA and rhizosphere microbiomes. However, more studies are warranted to understand the effect of HA fertilizer and beneficial microbiomes in responding to drought.

    • Climate change and soil erosion have provided unprecedented challenges to meet the rising demand for higher-quality food for the world's growing population. Over the past decades, maize productivity has experienced continuous improvement owing to technological innovations. The application of inorganic fertilizers has contributed to the intensity of agricultural production such as maize productivity[1]. The overuse of inorganic fertilizers, such as chemical nitrogen (N) fertilizer, can have adverse effects on both soil and groundwater[1]. Rather than relying on inorganic fertilizers derived from fossil fuels, using organic matter presents a promising alternative to remineralize depleted soil, enhance soil fertility, and ultimately increase crop productivity without adding to the environmental burden.

      Humic acid (HA) is an organic matter produced and accumulated by plants and animals. It is a residual substance through the decomposition of microorganisms accompanied by geochemistry processes[2]. A previous study suggested HA fertilizer could expand the effective absorption area of roots and enhance root growth[3]. Therefore, HA fertilizer promotes the recruitment of nutritional resources such as carbon (C) and N from both the fertilizer and bulk soil. Under water stress, studies showed that HA fertilization stimulates root development, i.e., increased root length and root dry weight, thereby delaying the drought stress responses[4]. However, the connections between HA and drought resistance, especially at the rhizosphere microbiome level, remain unclear.

      Recent evidence suggests that certain root-associated microbes likely mitigate the negative responses of drought stress, such as Actinobacteria and Firmicutes[5,6]. To test the hypothesis that HA delays negative drought responses in root development through enriching or depleting certain root-associated microbes, in this study, experiments were conducted to explore the effects of root-associated bacterial communities on several phenotypes during maize development and to determine how HA and drought influences the composition of root-associated microbial communities. Phenotypic traits for maize hybrids were collected during the flowering period under three different treatments (HA, drought, and control). The bacterial community was characterized from rhizospheres and bulk soils for the V4 region of the 16S rRNA gene using Illumina Hiseq2500 sequencer. A total of 18 samples (six individual plants per treatment) were collected for rhizosphere microbiome sampling, and six samples from the control were used for bulk soil microbiome sampling, resulting in ~763 million reads. Across all the samples, 40 bacterial phyla were detected in these four sample types (rhizosphere microbiomes from the control, HA, and drought group, as well as bacterial microbiomes of bulk soil from the control group).

    • The experiment was conducted on the Shandong Agricultural University research farm (Shandong, China). The soil was collected manually with sterile shovels from the farm and was then allocated equally into pots. Before planting, the soil was treated with commercial compound fertilizer (normal fertilization, N15-P15-K15) and commercial organic HA fertilizer (N15-P15-K15 + HA fertilization). Then, Xianyu335, a commercial maize hybrid, was planted with three treatments: (1) control group with normal fertilizer and conventional growth conditions; (2) HA group with HA fertilizer and conventional growth conditions; and (3) drought group with the normal fertilizer as that in the control group and moderate drought growth conditions (45% soil water content, SWC). The experiment was conducted following a completely randomized design with six replications per treatment.

    • The number of leaves (#Leaf) were counted on days 43, 49, 56, and 71 after sowing. Plant height (PH) was measured from the ground to the top of each plant, as described previously[7]. Days to silking (DTS) and days to pollen shed (DTP) were recorded for each plant.

    • Root samples were collected manually using a sterile shovel to a depth of approximately 20−30 cm, following a previously described method[8]. Sample collection was conducted during the flowering stage. Root samples were vortexed in sterile phosphate-buffered saline (PBS) buffer (Catalog No. E607008; Sangon Biotech, Shanghai, China) for 10 min at 14,000× g and centrifuged to obtain rhizosphere soil pellet after removing the root tissue. DNA extraction was performed using the FastDNA™ SPIN Kit for Soil (Catalog No. 116560200; FastDNA™, Solon, OH, USA) following the manufacturer's protocol. In brief, up to 500 mg of soil sample were added to a Lysing Matrix E tube with Sodium Phosphate Buffer, then were homogenized and centrifuged at 14,000× g for 10 min. The supernatant was transferred to a new microcentrifuge tube with a protein precipitation solution. After mixing, the tube was centrifuged at 14,000× g for 5 min. The supernatant was transferred and resuspended with binding matrix suspension to allow the binding of DNA. Finally, after quality checking the DNA by using a NanoDrop-2000 (Therml Fisher Scientific), the purified DNA was stored at −80 °C for further analysis.

    • After the quality checking of the DNA sample, all the qualified DNA was used to construct the 16S library. Briefly, the qualified DNA samples were amplified using a dual-indexed primer specific to the V4 region (515F (5'-GTGCCAGCMGCCGCGGTAA-3') and 806R (5'-GGACTACHVGGGTWTCTAAT-3')), and the PCR product was converted into blunt ends with T4 DNA polymerase, Klenow fragment, and T4 polynucleotide kinase. Then, after A tailing of the 3' end of each fragment, adapters were added. Then, AMPureXP beads (Beckman-Coulter, West Sacramento, CA, USA) was used to remove fragments that were too short. Finally, libraries quality checked by a Qubit 3.0 fluorometer (Life Technologies) were sequenced with paired-end 250-bp reads using Illumina Hiseq2500.

    • After sequencing, the raw data were cleaned by removing low-quality reads and reads contaminated by adapters, with a maximum of three bases mismatch allowed. The paired-end reads with overlap were merged into tags using FastLength Adjustment of Short reads (FLASH, v1.2.11)[9] with a minimal overlap length of 15 bp and the mismatching ratio of an overlapped region less than or equal to 0.1. Tags were then clustered into the operational taxonomic unit (OTU) using USEARCH (v7.0.1.1090)[10]. Principal component analysis (PCA) was then used to summarize factors mainly responsible for the differences of OTU composition in different samples, the similarity is high if two samples are closely located. Based on the OTU abundance information, the relative abundance of each OTU in each sample was be calculated, and the PCA of OTU was carried out with the relative abundance value. The Bray-Curtis distance matrix was used to assess the dissimilarity between samples. The software used in this step was package 'ade4' of the software[11] in R. To assess the statistical significance of the clustering observed in PCA, a PERMANOVA (Permutational Multivariate Analysis of Variance) was performed using the 'vegan' package[12] in R.

    • Student's t-test for phenotypic traits was used to determine the difference between different groups. The abundance differences of microbial communities between groups were detected by Wilcoxon signed-rank test and the p values were adjusted using the FDR approach[13]. Meanwhile, Linear Discriminant Analysis Effect Size (LEfSe) was incorporated to identify key taxa contributing to differences between conditions.

    • To investigate the potential effect of drought on plant growth and root-associated microbiomes, the plants were subjected to drought treatment (45% SWC). The rhizosphere and bulk soil samples were collected from maize hybrids to characterize microbial communities using next-generation sequencing (Fig. 1a). The phenotypic results revealed dramatic differences in plant height and root traits between control and drought conditions (Fig. 1b). The weights of both fresh (p < 0.01) and dry roots (p < 0.05) were significantly greater in the control group than drought group at the same stage (Fig. 1b bottom left, Fig. 1c & d). The plant height of the drought group was significantly shorter than that of the control group during the flowering stage (Fig. 1b middle left, Fig. 2a). Plant height was measured on days 43, 49, and 56 after planting. The results indicated that plant height at stages d49 and d56 was significantly decreased due to drought stress (Fig. 2a, p < 0.001). Meanwhile, the number of leaves were measured on days 43, 49, 56, and 71. The number of leaves were significantly higher in the control group at the earlier stage, however, more leaves were observed at the later stage in the drought group, and DTP were significantly delayed in the drought group (Fig. 2a, p < 0.05), which may indicate compensatory and delayed growth of plants due to drought stress.

      Figure 1. 

      Root performs significantly differently under drought stress vs normal conditions. (a) Flowchart of the study performed from the field work to the next generation sequencing. (b) The plant growth at days 42 (d42) and harvest at d72, as well as the root harvest at d72 under control and drought conditions, the scale bar is 20 cm. Statistics of the weight of (c) fresh roots, and (d) dry roots using R/Student's t test between each two of the control and drought groups. ** indicates p < 0.01, * indicates p < 0.05.

      Figure 2. 

      Summary statistics of phenotypes and relative abundance of the top 12 of the most abundant bacterial phyla. (a) Summary statistics of number of leaves (#Leaf) on days 43, 49, 56, and 71 after sowing, plant height (PH, cm) at days 43, 49, and 56 after sowing, and days to pollen shed (DTP, days) using Student's t test between cotrol and drought groups. * indicates p < 0.05, *** indicates p < 0.001. (b) Percent relative abundance of the most abundant phyla for control, drought treatments in rhizospheres and for control in bulk soils. All individuals were arranged in order within each group along the x axis. (c) Principal component analysis across the phyla from the Control Soil, control rhizosphere (Control Rhizo), and drought rhizosphere (Drought Rhizo) microbiome data.

    • A previous study on sorghum reported that drought delayed the development of the root microbiome and led to a higher abundance and greater activity of monoderm bacteria[13]. In this study, the effect of drought on root-associated microbial communities in maize was explored. As expected, the bulk soil microbiomes and maize rhizosphere microbiomes showed different patterns (Supplementary Fig. S1). The present results suggested that the composition of the rhizosphere at the phylum level caused by drought treatment differed significantly from that of the control (Figs 2b, 3; Supplementary Fig. S2). PCA was used to explore the differences in OTU composition across the samples. The PCA plot (Fig. 2c) showed a clear clustering of the samples based on their OTU profiles. Samples that were more similar in the drought group are located closer together, and similar patterns were observed in the control rhizosphere group, and the control soil group. The PERMANOVA results confirmed that the clustering observed in the PCA was statistically significant (p < 0.001). Consistent with the previous report on sorghum[13], the relative abundance of Actinobacterial and Firmicutes were significantly enriched and the relative abundance of the Proteobacterial and Fibrobacteres were significantly depleted in the rhizosphere microbiome of the drought group compared to the controls (Fig. 3a, p < 0.01). In addition, compared with the control group, some other microbes at the phylum level were significantly over-represented in the rhizosphere of the drought group, such as Euryarchaeota, Chlorobi, while some microbes at the phylum level were significantly depleted, such as Tenericutes (Fig. 3a, p < 0.01). LEfSe analysis was further conducted to identify key microbial taxa that were differentially abundant between the control and drought groups. These taxa, including Actinobacterial and Firmicutes, were visualized using a cladogram (Fig. 3b). The results indicated that these specific microbial groups were more prevalent under drought conditions, suggesting their potential role in shaping the microbial community structure in response to drought conditions.

      Figure 3. 

      (a) Wilcoxon Signed Rank Test for relative abundance of the abundant bacterial phyla, and (b) LEfSe analysis in control (red) and drought (darkgreen) groups. ** indicates p < 0.01.

    • Previous studies suggested that HA regulates plant growth by altering the root exudation profile[14] and played a role in protecting plants against water stress when co-inoculated with bacteria[15]. To further test the hypothesis that HA has a greater influence on the development of the root microbiome compared to the control during the flowering period, commercial compound fertilizers and organic HA fertilizer treatments were utilized. During the flowering stage, the HA group exhibited better-developed lateral roots compared to the control (Fig. 4a bottom left), although there were no statistically significant differences in both fresh and dry root weights between the two groups (Fig. 4b & c). Meanwhile, most of the phenotypes detected in this study showed minor differences between the control and HA group, except that the plant height at the earlier stage was significantly higher in the HA group (Supplementary Fig. S3). This result indicates that HA may play a potential role on belowground root traits but have little effect on aboveground traits.

      Figure 4. 

      Humic acid potentially benefits root growth vs normal conditions. (a) The plants grown on days 42 (d42) and harvested at d72, as well as the roots harvested on d72 under control and humic acid conditions, the scale bar = 20 cm. Statistics of the weight of (b) fresh roots, and (c) dry roots using R/Student's t test between the control and humic acid conditions. 'NS' indicates not significant.

      To better understand the potential role of HA fertilization on the root microbiome, the root microbiome under HA fertilization was determined. Phylum-level relative abundance of Acidobacteria, Acitinobacteria, Bacteroidetes, Crenarchaeota, Cyanobacteria, and Proterobacteria revealed that the rhizosphere microbiomes exhibited a different composition compared to the control (Fig. 5a, 'Control Rhizo' and 'Humic Acid Rhizo'; results of statistical analyses were shown in Supplementary Fig. S4). While the above results suggest a different pattern for an abundance of these rhizosphere microbiomes on the phylum level during the development of the root microbiome under HA-treated environments. Evidence of the root rhizosphere microbiome is observable in these microbial lineages when a finer taxonomic resolution was used, as illustrated by family-level changes (Fig. 5b; results of statistical analyses are shown in Supplementary Fig. S5).

      Figure 5. 

      Relative abundance of the top 12 of the most abundant bacterial phyla and families. Percent relative abundance of the (a) most abundant phyla, and (b) family for (c) control, humic acid (H) treatments in rhizospheres and for control in bulk soils (S). All individuals were arranged in order within each group along the x axis.

    • This study identified 40 bacteria phyla across soil, control rhizosphere, drought-induced, and HA-induced rhizosphere bacteria. The diversity of 12 out of 40 bacterial communities at the phyla level significantly varied in their resistance to drought stress (Supplementary Fig. S2). Among these, Actinobacteria and Bacteroidetes were notably enriched and depleted, respectively, in HA fertilizer-treated samples (Supplementary Fig. S4), which was consistent with a previous study[16]. In this study, the increased abundance of Actinobacteria might be caused by the HA increase in the N content in the soil, further enhancing the rate of nitrogen cycling[16]. Actiomyces are known for their ability to produce enzymes that degrade complex organic matter, which could enhance the availability of nutrient uptake. This nutrient enrichment might synergize with HA, thus improving plant growth and resistance under stress conditions. While the present study primarily focused on the most abundant bacterial phyla and families, it is crucial to consider the role of low-abundant yet core microbial taxa. These taxa, although present at lower relative abundances, often contribute significantly to ecosystem stability and plant health. Low-abundance microbial communities can serve as keystone species, providing essential functions such as nutrient cycling, production of growth-promoting hormones, and suppression of pathogenic organisms. Targeted approaches in further studies, such as metagenomics or metatranscriptomics, could help elucidate the specific roles of these low-abundant microbial taxa in shaping plant phenotypes and contributing to overall plant fitness under varying environmental conditions.

      To improve maize drought resistance and harness the benefits of HA for agricultural productivity, manipulating the root microbiome provides a promising opportunity. However, it is essential to understand the causes and consequences of shifts in the soil and root microbiome induced by drought and HA. In the current study, drought delayed plant development, measured by the number of leaves and plant height. However, as plants under normal conditions began reproductive growth and experience senescence, the drought-treated plants showed a significantly higher leaf count, consistent with previous observations[17]. Drought conditions were found to alter the microbial diversity, leading to an increase in drought-resistant phyla such as Actinobacteria and Firmicutes and a decrease in drought-sensitive phyla such as Bacteroidetes and Proteobacteria (Supplementary Fig. S2). These shifts could significantly impact crop performance by influencing nutrient availability and stress resistance. Understanding these microbial shifts could provide valuable insights into how the microbiome could contribute to adaptation to environmental changes. A previous study on physiological and molecular mechanisms of HA influencing drought resistance in maize, demonstrated that the application of HA under drought conditions improved the proportion of soil macro-aggregates (e.g., P, K, Fe, and Mg), increased ATP synthase activity, enhanced the content of IAA, and the concentration of osmotically active solutes[18]. This previous study also showed that maize yields were improved with HA treatments under drought stress. However, how HA fertilization impacts belowground root traits and microbiome diversity remains largely unknown. In this study, HA demonstrated a potential positive effect on root architecture, leading to more extensive root development. However, it is important to understand why these changes in root structure did not translate to statistically significant differences in root biomass. A possible reason may be due to the physiological mechanisms influenced by HA. Previous studies[4,18] suggest that HA can enhance root elongation and branching without necessarily increasing total root weight. The improvements in root architecture may be related to changes in root surface area, which facilitates nutrient and water uptake, rather than an increase in root mass. This improvement may not significantly affect overall root weight within the short-term duration of the present study.

      The current study did not perform the HA treatment under drought stress. Whether these potentially beneficial microbiomes will be recruited with HA treatments under drought conditions needs to be tested with additional experiments. Nevertheless, the results indicate that HA affects the abundance and composition of rhizosphere microbiome and underscore the importance of root-associated microbiomes in responding to drought stress (Supplementary Fig. S6).

      • We would like to thank Dr. Fupeng Song for assistance in soil water content determination. We appreciate the constructive suggestions from anonymous reviewers. This study was funded by the National Key Research and Development Program of China to H. Liu (2022YFD1201700), the National Natural Science Foundation of China to X. Yang (32101705), and the Youth Innovation team project to Xuerong Yang.

      • The authors confirm contribution to the paper as follows: study conception and design: Yang X; experiment conduction: Sun M, Zhang L, Kang C, Li X, Sun Y, Zeng X, Dong L; data analysis: Yang X, Zhu H, Liu H; draft manuscript preparation: Yang X, Liu H. All authors critically revised and provided final approval of this manuscript.

      • The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.

      • The authors declare that they have no conflict of interest.

      • # Authors contributed equally: Mingfei Sun, Lin Zhang

      • Supplementary Fig. S1 Heatmap of the bacteria phylum (A) as well as Shannon’s diversity (B&C) represented in the control soil (S), control (C), drought (D) and HA (H) rhizosphere microbiome.
      • Supplementary Fig. S2 Wilcoxon Signed Rank Test for percent relative abundance of the abundant bacterial phylum in Control Soil, Control rhizosphere (Control Rhizo) and drought rhizosphere (Drought Rhizo) groups.
      • Supplementary Fig. S3 Summary statistics of phenotypes.
      • Supplementary Fig. S4 Wilcoxon Signed Rank Test for relative abundance of the abundant bacterial phyla in control soil, control rhizosphere and Humic Acid rhizosphere groups.
      • Supplementary Fig. S5 Summary statistics of relative abundance of the top 12 of the most abundant baterial families.
      • Supplementary Fig. S6 Summary analyses of LEfSe among control soil (S), control (C), drought (D) and HA (H) rhizosphere conditions.
      • Copyright: © 2024 by the author(s). Published by Maximum Academic Press, Fayetteville, GA. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
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    Sun M, Zhang L, Zhu H, Kang C, Li X, et al. 2024. Responses of microbial communities to drought stress and humic acid fertilizer in maize. Genomics Communications 1: e006 doi: 10.48130/gcomm-0024-0006
    Sun M, Zhang L, Zhu H, Kang C, Li X, et al. 2024. Responses of microbial communities to drought stress and humic acid fertilizer in maize. Genomics Communications 1: e006 doi: 10.48130/gcomm-0024-0006

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