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Article

Miscanthus × giganteus Rhizobacterial Community Responses to Zn and Oil Sludge Co-Contamination

by
Asil Nurzhanova
1,
Eugenia Boulygina
2,
Irina Sungurtseva
3,
Aigerim Mamirova
4,*,
Ramza Berzhanova
4 and
Anna Muratova
3
1
Institute of Plant Biology and Biotechnology, Timiryazev 45, Almaty 050040, Kazakhstan
2
Institute of Fundamental Medicine and Biology, Kazan Federal University, Marx, 76, 420012 Kazan, Russia
3
Institute of Biochemistry and Physiology of Plants and Microorganisms, Saratov Scientific Centre of the Russian Academy of Sciences (IBPPM RAS), Entuziastov 13, 410049 Saratov, Russia
4
Department of Biotechnology, Faculty of Biology and Biotechnology, Al-Farabi Kazakh National University, Al-Farabi 71, Almaty 050040, Kazakhstan
*
Author to whom correspondence should be addressed.
Agronomy 2025, 15(9), 2232; https://doi.org/10.3390/agronomy15092232
Submission received: 21 August 2025 / Revised: 17 September 2025 / Accepted: 19 September 2025 / Published: 22 September 2025
(This article belongs to the Special Issue Innovations and Obstacles: Microbial Communities in the Journey of Soil Remediation)
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Abstract

Soil contamination in industrial areas often involves complex mixtures of contaminants, making remediation a significant challenge. Microbe-assisted phytoremediation offers a promising solution, yet its success depends on understanding interaction between plants, microorganisms, and contaminants in rhizosphere. This study examined the effects of organic (oil sludge) and inorganic (Zn) contaminants, applied individually and in combination, on the rhizosphere bacterial community of Miscanthus × giganteus Greef et Deu (M×g), with emphasis on strains exhibiting plant growth-promoting, hydrocarbon-degrading, and metal-tolerant traits. A one-season greenhouse experiment included soils spiked with Zn (1650 mg kg−1) and/or oil sludge (15 mL kg−1). Oil sludge exerted a stronger influence on the taxonomic structure of rhizobacterial communities than Zn, largely shaping the patterns observed under co-contamination. Zn exposure increased the relative abundance of Actinobacteriota, whereas oil sludge favoured Proteobacteriota. Both contaminants, individually and together, enhanced the proportion of Sphingomonadaceae. Across all treatments, taxa with potential plant-growth-promoting traits were present, while co-contaminated soil harboured microorganisms capable of hydrocarbon degradation, heavy metal tolerance, and plant growth promotion. These findings highlight the adaptive capacity of the M×g rhizobiome and support its application in phytoremediation. The isolation and characterisation of rhizosphere-associated strains provide basis for developing microbial bioagents to enhance biomass production and remediation efficiency in multi-contaminated environments.

1. Introduction

Under real conditions, technogenic soil pollution typically involves complex mixtures of contaminants, with petroleum hydrocarbons and heavy metals being the most prevalent [1,2,3,4,5,6]. A recent study estimated that 14–17% of cropland soils already exceed agricultural thresholds for at least one heavy metal [7]. Hydrocarbons are classified as priority pollutants because of their persistence, high toxicity, mutagenic and carcinogenic potential, and teratogenic effects in humans [8]. Combined contamination commonly occurs in industrial zones of oil extraction and processing, roadside soils, and landfill areas [4,5,9,10,11]. The coexistence of pollutants with distinct physicochemical properties poses a significant challenge for remediation, as they respond differently to conventional rehabilitation strategies [9]. In this context, phytoremediation, which relies on vegetation to restore contaminated soils, is regarded as one of the most effective modern technologies, as it enables the simultaneous removal of both organic and inorganic pollutants through diverse mechanisms [12]. The principal mechanism for the remediation of organic pollutants, such as petroleum hydrocarbons, is rhizosphere-mediated microbial degradation [13,14], whereas heavy metal remediation primarily relies on phytoaccumulation and phytostabilisation [15,16,17]. The efficiency of these processes is determined by both the remediating plant and its associated rhizosphere microbiome.
In recent years, plants of the Miscanthus genus have attracted considerable interest owing to their economic value and broad applicability across multiple sectors [18,19]. Among these, Miscanthus × giganteus Greef et Deu (M×g) is particularly recognised as a promising bioenergy crop capable of establishing in marginal and abandoned lands. This species is characterised by high biomass productivity and has shown potential for the remediation of soils contaminated with both organic and inorganic pollutants [20,21,22,23,24,25]. The ability of plants to diverse environmental conditions is strongly influenced by ecological services provided by their microbial symbionts, including biofertilisation, disease suppression, and enhanced tolerance to abiotic stress [26]. Soil functions as a reservoir of microbial diversity from which plants selectively assemble their microbiomes according to the requirements of their specific habitats [26]. Understanding the formation of rhizosphere microbial communities that support plant growth under adverse conditions and facilitate soil remediation is essential for evaluating the feasibility of cultivating M×g in contaminated environments.
The microbial communities associated with M×g have been extensively studied using both culture-independent and culture-based approaches. Since the majority of microorganisms are non-culturable (>99%), even when diverse media are employed [27,28], modern culture-independent molecular techniques are indispensable for assessing microbial diversity. A comprehensive study of 26 plant species identified Clavibacter, Chryseobacterium, and Acidovorax as taxa uniquely associated with M×g [29]. Increasing evidence indicates that Miscanthus species actively shape their rhizosphere microbiomes to support growth in disturbed soils [30,31,32,33,34]. Predictive functional profiling using Phylogenetic Investigation of Communities by Reconstruction of Unobserved States (PICRUSt) revealed that Miscanthus cultivation modifies the metabolic capabilities of bacterial communities, including pathways related to carbon and nitrogen metabolism and the phosphorus cycle [35]. Furthermore, FUNGuild analysis demonstrated that Miscanthus cultivation alters the trophic modes of fungal communities [35]. Miscanthus-associated diazotrophs supply plants with bioavailable nitrogen [36,37,38], and nitrogen fertilisation has been shown to influence the composition of the Miscanthus microbiome [39]. Several factors affecting the assembly of microbial communities in the Miscanthus rhizosphere are currently under investigation, including plant species identity [40], genetic variability [41], plant age [39], rooting depth [42], and exposure to different pollutants [22,43,44,45,46].
Several studies have documented shifts in the composition and structure of the M×g rhizosphere microbial community in response to heavy metal contamination [22,44,46,47]. In contrast, information on the effects of hydrocarbons on the M×g rhizosphere microbiome remains scarce [48]. These observations provide a basis for further investigations into the ecological functions of microorganisms in Miscanthus growth and offer valuable insights for the cultivation of this species in marginal soils, particularly those contaminated with heavy metals or other pollutants.
Increasing attention has recently been directed toward understanding plant responses and remediation efficiency in soils contaminated with multiple pollutants, particularly heavy metals and hydrocarbons [49,50,51,52]. However, only three studies published in 2025 have investigated the effects of complex pollution on the root zone microbiome. Conlon et al. [53] examined rhizodegradation by Sinapis alba, Lolium perenne, L. perenne + Trifolium repens, and Cichorium intybus in petroleum hydrocarbon- and heavy metal-co-contaminated soil in a microcosm-scale pot trial, reporting rhizosphere enrichment with hydrocarbon-degrading bacteria, including KCM-B-112, C1-B045, Hydrogenophaga, unclassified Saccharimonadales sp., and Pedobacter. Another microcosm experiment demonstrated that elevated levels of Cu, Ni, and Zn (100 mg kg−1 each) negatively affected the growth of Lolium multiflorum and reduced the abundance of hydrocarbon-degraders, while also altering the structure of hydrocarbon-degrading bacterial communities [54]. In a field study evidencing the co-contaminated nature of soils in industrial areas, the rhizosphere of Carmona microphylla, a native plant growing at an operating ink factory site polluted with polycyclic aromatic hydrocarbons (~500–800 mg kg−1) and heavy metals (As, Cd, Cr, Pb and Zn), was analysed [55]. High pollution levels markedly reduced the number of specific operational taxonomic units (OTUs), with heavy metals identified as the primary factor shaping bacterial community structure [55]. Soil bacterial functions were influenced mainly by pollution levels rather than rhizosphere effects, with high contamination altering α diversity as well as the structure and composition of C- and N-cycling bacteria. Compared to hydrocarbons, heavy metals played a more significant role in shaping bacterial communities by affecting metabolite profiles [55]. Our previous research described the physiological and biochemical responses of M×g to soils contaminated with heavy metals and petroleum hydrocarbons, as well as its effective application in remediating soils affected by mixed pollution [56]. Zinc (Zn) and oil sludge, both widespread anthropogenic contaminants [8,57], were selected as model pollutants in the present study. Elevated concentrations of these pollutants in soils often result from industrial activities such as mining and smelting [10].
This study investigated the rhizobacterial community of M×g cultivated in soils subjected to distinct contamination scenarios: Zn, oil sludge, or their combination, with uncontaminated soil serving as a control. The research was grounded in the premise that soil contamination adversely affects rhizosphere microbial diversity, which is essential for both plant health and soil functioning [58,59]. It was hypothesized that complex contamination involving both inorganic (Zn) and organic (oil sludge) pollutants would exert a more pronounced negative effect on rhizosphere microbial diversity than single-pollutant exposure. In particular, a decline was expected in the abundance and diversity of cultivable microbial taxa with plant growth-promoting traits and contaminant tolerance [54]. Moreover, it was anticipated that organic contamination (oil sludge) would be less detrimental, or potentially more favourable, to rhizobacterial community composition than inorganic contamination (Zn), owing to differences in bioavailability, toxicity mechanisms, and the potential for microbial degradation. Within this framework, the study aimed to elucidate how contamination type and complexity shape rhizosphere microbial communities and their functional potential in the context of phytoremediation with M×g.

2. Materials and Methods

2.1. Soil Origin and Characteristics

The experimental design was previously described by Muratova et al. [56]. Briefly, the soil used in the pot experiments was loamy leached chernozem (World Reference Base [60]: Luvic Chernozems) collected near Novye Burasy village, Saratov region, Russian Federation. Prior to the experiment, the soil was analysed for agrochemical parameters. Soil pH was measured using a Metler Toledo Delta 320 pH meter (Mettler-Toledo Instrument Co., Ltd.; Shanghai, China) [61]. The contents of mobile (available) phosphorus (P2O5) [62], nitrate (NO3-N) [63], and water-soluble ammonium (NH4-N) [64] were quantified by photocolorimetry (Table 1). The principal characteristics of the soil were as follows: particle-size distributions—1–3 mm, 0.036%; 1–0.5 mm, 0.23%; 0.5–0.25 mm, 31.1%; and <0.25 mm, 68.5%. The total organic carbon content was 4.4% of the air-dried soil.

2.2. Experimental Layout

The soil for the pot experiment was sieved through a 7 mm mesh and divided into four portions. The first portion was spiked with a Zn solution obtained by dissolving ZnSO4 · 7H2O (>99% purity, Reakhim, Russia) in deionised water, resulting in a final soil concentration of 1650 mg Zn kg−1, corresponding to a 15 × MPC [65]. The second portion was spiked with oil sludge (15 mL kg−1), yielding a final concentration of total petroleum hydrocarbons (TPHs) of 8810 mg kg−1. The oil sludge was collected from a pit located on the grounds of a petroleum refinery in Saratov, Russian Federation. The oil sludge was collected from an oil sludge pit located on the grounds of a petroleum refinery in Saratov, Russian Federation. Its characteristics were as follows: pH 6.4, suspended solids ≤ 2%, water content 14.2%. The petroleum fraction comprised paraffins (18.8%), naphthene (18.1%), mono- and bicyclic aromatic hydrocarbons (MBCA, 7.1%), polycyclic aromatic hydrocarbons (PAHs, 11.2%), and alcohol-benzene tars (ABTs) plus asphaltenes (44.7%). The third portion was spiked with both the Zn solution and oil sludge at the same final concentrations. The fourth portion served as the control and was treated with an equivalent amount of distilled water. Thus, the experimental treatments were: (i) control soil, (ii) Zn-spiked soil, (iii) oil sludge-spiked soil, and (iv) Zn- and oil sludge-spiked soil.
Pots were lined with black plastic bags to prevent leaching of soil solution, including water-soluble contaminants and nutrients. A 1 kg layer of expanded clay (granule diameter: 15–20 mm) was placed at the bottom of each pot as drainage and covered with gauze. This was overlaid with 1 kg of purified river sand, which was again covered with gauze. Finally, 3 kg dry weight (DW) of either control or spiked soil was added. Two one-year-old M×g rhizomes (sourced from the plantation of the Institute of Plant Biology and Biotechnology, Almaty, Kazakhstan) were planted in each pot, and the soil surface was covered with 200 g of purified river sand.
All treatments were established in triplicate. Soil moisture was maintained at 50% of field moisture capacity by daily watering, determined gravimetrically. Plants were cultivated in a greenhouse under natural illumination at 24–28 °C during the day and 20–22 °C at night for six months (March–September 2018).

2.3. Isolation of Strains Capable of Tolerating Contaminants and Promoting Plant Growth

Heavy-metal-resistant microorganisms were enumerated and isolated using Lysogeny Broth (LB) agar supplemented with either ZnSO4 · 7H2O or Pb(CH3COO)2 at a final metal-ion concentration of 0.5 mmol L−1. The isolates were subsequently tested for metal resistance by culturing on LB agar containing 0.5–5 mM Zn2+ or Pb2+.
All isolates were assessed for plant growth–promoting traits, including the ability to fix atmospheric nitrogen (N2) on nitrogen-free agar medium [66,67], produce the phytohormone indole-acetic acid (IAA) [68], and solubilise phosphates [69]. The paraffin-degrading ability of isolates was tested using the membrane-filter technique and Bushnell–Haas medium [67]. PAH degradation was evaluated by a plate assay based on the formation of clearance zones around bacterial colonies [70].

2.4. 16S rRNA Gene Sequencing Analysis of Rhizosphere Soil

At the end of the experiment, plant roots were removed from the pots and soil loosely adhering to the roots was shaken off. Young roots with tightly attached rhizosphere soil were collected (~1.0 g) for metagenomic analysis. Prior to analysis, rhizosphere soil was carefully separated from the roots using a laboratory spatula. Two replicate subsamples (0.3 g each) were taken for DNA extraction.
Soil DNA was extracted and purified using the Fast DNA SPIN Kit for Soil (MP Biomedicals, Solon, OH, USA) and homogenised with a FastPrep-24 instrument (MP Biomedicals, Solon, OH, USA) according to the manufacturer’s instructions. A 16S rRNA gene sequencing library was constructed following the Illumina 16S Metagenomics Sequencing Library Preparation protocol (Illumina, San Diego, CA, USA), targeting the V3–V4 hypervariable regions. The initial PCR amplification was carried out with region-specific primers containing Illumina adapter overhang sequences (forward primer: 5′-TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGCCTACGGGNGGCWGCAG-3′; reverse primer: 5′-GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGGACTACHVGGGTATCTAATCC-3′). PCR products were visualised by gel electrophoresis and purified with AMPure XT magnetic beads. A second PCR was then performed using the Nextera XT Index Kit (Illumina, San Diego, CA, USA).
The DNA concentration of purified PCR products was quantified with a Qubit dsDNA HS Assay Kit (Thermo Scientific, Waltham, MA, USA) on a Qubit 2.0 fluorometer. The pooled library (4 nM) was denatured with 0.2 M NaOH, diluted to 10 pM, and combined with 20% (v/v) denatured 4 pM PhiX control, according to Illumina’s guidelines. Sequencing of the V3–V4 regions was performed on an Illumina MiSeq platform using a 2 × 300 bp paired-end run.
Raw sequence reads were deposited in the NCBI Sequence Read Archive (SRA) under BioProject accession number PRJNA1119175 in FASTQ format (https://www.ncbi.nlm.nih.gov/bioproject/PRJNA1119175, accessed on 14 August 2025).

2.5. Bioinformatics and Statistical Analysis

Sequence data were processed using QIIME2 software, version 2022.8 (http://qiime2.org/; accessed on 1 August 2025) [71]. On average, 118,339 read pairs per sample were obtained before filtering. Raw reads were denoised with the DADA2 algorithm implemented in QIIME2 [72]. After quality filtering and removal of chimeric and phiX sequences, an average of 17,870 joined read pairs per sample remained. Taxonomic classification was performed using the SILVA 138 database (99% OTUs) and a Naive Bayes classifier [73]. The number of observed features ranged from 601 to 1030.
Alpha (α) diversity, reflecting richness and evenness of the bacterial community, was estimated by using Chao1, Shannon, and Simpson indices. Beta (β) diversity, representing differences in microbial community composition among samples, was assessed using weighted and unweighted UniFrac distances and visualised by non-metric multidimensional scaling (nMDS). Venn diagrams were constructed with Creately software (https://creately.com/lp/venn-diagram-maker/; accessed on 1 August 2025).
Statistical analyses were performed with Statistica 13 (TIBCO Software Inc. 2017, Palo Alto, CA, USA). Fisher’s least significant difference (LSD) test was used to evaluate treatment effects. Mean comparisons were conducted with Tukey’s HSD, and Spearman’s rank correlation coefficients were also calculated.

3. Results

3.1. Diversity of Rhizosphere Communities

Sequencing of the 16S rRNA gene from 10 rhizosphere samples generated 1,122,256 raw reads. After denoising and chimera removal, 703,550 sequences were retained for downstream analyses. Sequences with >97% similarity were clustered into OTUs, which were assigned to 32 phyla, 98 classes, 229 orders, 346 families, and 587 genera. Rarefaction curves based on normalised OTU counts nearly reached saturation for all samples, indicating sufficient sequencing depth and comprehensive coverage of the bacterial communities (Figure S1).
Bacterial diversity and richness were evaluated using α- and β-diversity metrics (Table 2, Figure 1).
The α-diversity of rhizosphere bacterial communities was assessed using species richness indices (Chao1, Shannon, and Simpson) and Faith’s phylogenetic diversity (Table 2). The rhizobacterial community of M×g grown in control soil exhibited the greatest richness, as indicated by the observed features and Chao1 indices. Rhizobacterial community of plants grown in Zn-spiked and oil sludge-spiked soils showed significantly reduced richness (observed features: p = 0.04; Chao1: p = 0.007). The lowest overall richness, including phylogenetic diversity (Faith PD, p = 0.008), was recorded in oil sludge-spiked soil. By comparison, the highest diversity was detected in rhizobacterial communities from plants grown in Zn- and oil sludge-co-contaminated soil, where both Shannon and Simpson indices reached their maximum values.
Non-metric multidimensional scaling (nMDS) visualisation further demonstrated that soil treatment significantly shaped the composition of M×g rhizosphere communities, with clear clustering of samples according to treatment (Figure 1).

3.2. Taxonomic Structure of Rhizosphere Communities

MiSeq sequencing revealed that the M×g rhizosphere communities comprised 587 bacterial genera belonging to 346 families across 32 phyla. The predominant bacterial phyla under different contamination conditions are shown in Figure 2.
The rhizosphere bacterial community of M×g grown in control loamy leached chernozem was dominated by bacteria rather than archaea. Across all samples, the dominant phyla were Pseudomonadota (13 ± 2.5% to 39 ± 4.1%), Actinobacteriota (12 ± 0.1% to 27 ± 5.5%), and Firmicutes (13 ± 0.8% to 27 ± 0.6%). However, their relative abundances varied depending on the type of contaminant. A high proportion of Firmicutes in the control soil was associated with the use of material stored without moistening for one year. Contamination with both heavy metal and hydrocarbons reduced the abundance of Firmicutes, although this group remained dominant.
Statistical analysis confirmed that soil treatments significantly influenced the composition of M×g rhizosphere communities (Figure 1). The addition of contaminants decreased the proportion of Firmicutes, with oil sludge exerting a stronger effect than Zn (Figure 2). Zn contamination significantly (p < 0.05) increased the proportions of Actinobacteriota (by 54%), Pseudomonadota (by 37%), and Bacteroidota (by 69%), while decreasing those of Firmicutes (by 45%), Planctomycetota (by 51%), and Acidobacteriota (by 20%). Conversely, oil sludge contamination promoted the growth of Pseudomonadota (by 186%), Acidobacteriota (by 84%), and Verrucomicrobia (by 56%), while reducing the abundance of Actinobacteriota (by 41%), Bacteroidota (by 46%), Chloroflexota (by 51%), Firmicutes (by 52%), and Gemmatimonadetes (by 54%). The taxonomic profile of communities under mixed contamination was intermediate, reflecting the combined but distinct impacts of each contaminant.
Family-level taxa identified in control, Zn-spiked, oil sludge-spiked, and Zn- and oil sludge-co-spiked soils are listed in Table S1. Statistically significant differences were observed in the community composition of oil sludge-spiked soil compared with Zn-spiked soil (LSD = 0.025299), control soil (LSD = 0.021431), and co-contaminated soil (LSD = 0.030404).
Quantitative characteristics of predominant taxa (≥0.5%) identified at the genus level, including members with potential roles as Zn and oil sludge detoxifiers and/or plant growth promoters, are shown in Figure 3.
Among the Actinobacteriota, the dominant orders in the M×g rhizosphere were Micrococcales and Gaiellales, each maintaining a relative abundance of ~4% across all treatments. The predominant families included Micrococcaceae, Micromonosporaceae, Rubrobacteriaceae, Gaiellaceae, and an uncultivated family within Gaiellales. Zn contamination promoted Actinobacteriota growth, showing strong correlations with increased proportions of Intrasporangiaceae (2.4-fold; R2 = 0.830, p < 0.05) and Micrococcaceae (6.4-fold; R2 = 0.976, p < 0.05). Enrichment of the rhizosphere with Pseudarthrobacter (Micrococcaceae), Micromonospora (Micromonosporaceae), and Nocardioides (Nocardioidaceae) was also observed, with increases of 5.6-, 5.5-, and 4.3-fold, respectively (Figure 3).
In contrast, oil sludge reduced the abundance of several Actinobacteriota families. In oil sludge-spiked soil, Intrasporangiaceae, Micrococcaceae, Rubrobacteriaceae, Micromonosporaceae, Nocardioidaceae, and Propionibacteriaceae decreased by 2.0-, 2.0-, 3.9-, 5.8-, 2.8-, and 6.5-fold, respectively, compared with the control (Table S1). Strong correlations were detected between oil sludge presence and increased proportions of OTUs affiliated with Nocardiaceae (R2 = 0.995, p < 0.05) and Microbacteriaceae (R2 = 0.959, p < 0.05). Oil sludge also promoted the growth of Microbacterium and Mycobacterium (Figure 3).
The effects of oil sludge, both alone and in combination with Zn, significantly increased the proportions of families within the phylum Proteobacteriota, particularly those belonging to Alphaproteobacteria. In oil sludge-spiked soils, Alphaproteobacteria families were enriched as follows: Sphingomonadaceae (3.4-fold), Xanthobacteraceae (1.3-fold), Micropepsaceae (3.8-fold), Rhizobiaceae (2.2-fold), and Parvibaculaceae (2.2-fold) (R2 = 0.990, p < 0.05). Betaproteobacteria also increased, with Burkholderiaceae (13-fold), Oxalobacteraceae (1.7-fold), and Comamonadaceae (1.4-fold) showing enrichment (R2 = 0.955, p < 0.05). Within Gammaproteobacteria, increases were observed in Acidithiobacillaceae, Porticoccaceae, and Xanthomonadaceae (R2 = 0.950, p < 0.05), compared to the control.
At the genus level, the most notable increases under oil sludge contamination included KCM-B-112 (Acidithiobacillaceae), Sphingomonas (Sphingomonadaceae), uncultured Micropepsaceae, Parvibaculum (Parvibaculaceae), Acidibacter, Noviherbaspirillum (Oxalobacteraceae), SC-I-84, C1-B045 (Porticoccaceae), Immundisolibacter (Immundisolibacteraceae), and Lysobacter (Xanthomonadaceae) (Figure 3). Zn contamination also significantly increased the abundance of Micropepsaceae (1.3-fold), Sphingomonadaceae (1.9-fold), Burkholderiaceae (7.5-fold), and Rhodanobacteraceae (2.0-fold) compared to the control.
Genera enriched under Zn treatment included Sphingomonas (Sphingomonadaceae), Aquicella (Diplorickettsiaceae), SC-I-84 (Burkholderiales order), Methylobacterium-Methylorubrum (Beijerinckiaceae), and Phenylobacterium (Caulobacteraceae) (Figure 3). Under mixed contamination, the proportions of Rhizobiaceae, Sphingomonadaceae, Burkholderiaceae, and Rhodanobacteraceae were significantly higher (R2 = 0.999, p < 0.05), while Nitrosomonadaceae (SC-I-84 genus) was reduced compared with the other treatments. Genera enriched in Zn- and oil sludge-co-contaminated soils included Sphingomonas (Sphingomonadaceae), KCM-B-112 (Acidithiobacillaceae), Burkholderia-Caballeronia-Paraburkholderia (Burkholderiaceae), Rhodanobacter (Rhodanobacteraceae), Mesorhizobium (Rhizobiaceae), Pseudomonas (Pseudomonadaceae), and Legionella (Legionellaceae) (Figure 3).
Oil sludge also strongly promoted Acidobacteriota. The abundance of Koribacteraceae increased by 838% (R2 = 0.984), while uncultivated Acidobacteriales increased by 154% (R2 = 0.969). In contrast, Zn, either alone or in combination with oil sludge, did not significantly affect Acidobacteriota (Table S1). Within this phylum, the genera Bryobacter, Candidatus_Koribacter, and Candidatus_Solibacter were significantly enriched in oil sludge-spiked soil (Figure 3).
Within the Bacteroidota phylum, oil sludge altered community structure by greatly increasing Sphingobacteriaceae (19-fold under oil sludge, 25-fold under Zn + oil sludge), with Mucilaginibacter emerging only under oil sludge (Figure 3). In contrast, Chitinophagaceae decreased (4.3-fold under oil sludge, 1.4-fold under Zn + oil sludge), with Flavisolibacter disappearing under oil sludge exposure (Figure 3). Zn contamination alone did not significantly affect Bacteroidota composition.
The Chloroflexota phylum showed marked reductions in predominant families under oil sludge contamination, while Zn contamination had no significant effect. Oil sludge also reduced Bacillaceae abundance in the M×g rhizobacterial community by twofold, with similar reductions under Zn and mixed contamination. Bacillaceae abundance was closely correlated with the presence of oil sludge (R2 = 0.953, p < 0.05). In addition, oil sludge reduced Gemmatimonadaceae abundance by 2.2-fold, with a further 3.4-fold decrease under Zn + oil sludge co-contamination. Both Zn and mixed contamination supressed members of the Planctomycetota phylum, significantly reducing the predominant unspecified family of the WD2101 order by 4.0- and 2.5-fold, respectively (Figure 3).

3.3. Common and Unique Taxa Among Rhizosphere Bacterial Communities

Comparative analyses were performed to identify OTUs common or specific to the rhizosphere of M×g under control contamination treatments (Figure 4). A total of 126 genera were shared across all conditions, representing approximately one-third of the rhizosphere communities.
Control and mixed-contaminated soils exhibited the highest total number of OTUs at the genus level, as well as the largest number of unique genera (Figure 4). In control soil, only 38.1% of genera were shared with other treatments, while 18.4% (61 genera) were unique. These taxa were mainly affiliated with Firmicutes (9 genera), Gammaproteobacteria (8), Verrucomicrobiota (7), Myxococcota (6), Alphaproteobacteria (6), and Patescibacteria (5) (Table S2).
The Zn-spiked rhizosphere soil shared 40.8% of genera with other treatments, with 14.2% (44 genera) unique. Unique taxa in this soil were primarily affiliated with Firmicutes (9 genera), Bacteroidota (6), Alphaproteobacteria (6), Actinobacteriota (5), and Patescibacteria (5) (Table S3).
Oil sludge-spiked rhizosphere soil contained 47.4% of genera in common with other treatments, with the lowest proportion of unique taxa (12.0%; 32 genera). Unique genera were mainly Alphaproteobacteria (11 genera), with no unique Firmicutes detected (Table S4).
Rhizosphere soil co-contaminated with Zn and oil sludge had the highest total number of genera (344), of which only 36.6% were shared with other treatments. This community also contained the largest proportion of unique taxa (20.6%; 71 genera; Table S5). These were predominantly Gammaproteobacteria (20 genera), Actinobacteriota (12), Firmicutes (12), and Bacteroidota (6).

3.4. Plant-Growth Promoting and Contaminant-Tolerant Microbial Strain Isolation

From polluted rhizosphere samples, 76 microbial strains were isolated and screened for heavy metal resistance and plant growth-promoting traits, including nitrogen fixation, phosphorous solubilisation, and production of siderophores and phytohormones. Of these, 12 isolates were resistant to Pb2+ or Zn2+ at concentrations >5 mM, 10 isolates degraded paraffins, and 2 degraded phenanthrene. In addition, 33 isolates synthesised the phytohormone IAA, 10 fixed atmospheric N2, 8 produced siderophores, and 2 solubilised phosphorus. Isolates combining plant-growth-promoting and pollutant-resistance traits were selected, identified, and are listed in Table 3.

4. Discussion

The ability of M×g to colonise and grow on marginal, disturbed, and polluted lands without compromising biomass accumulation has attracted considerable attention [74]. Such adaptability is determined not only by plant genetics but also by the microorganisms associated with its roots. Soil microorganisms are known to induce plant resistance to adverse environmental conditions, including drought and anthropogenic pollution [75,76]. Plants selectively recruit microorganisms from surrounding soil that enhance their survival, making root-associated microbiota crucial for successful colonisation and establishment [26]. Among these microorganisms, bacteria represent the most abundant group [77]. Accordingly, this study investigated how bacterial community composition in the rhizosphere of M×g is shaped by different types of soil contamination.
Our comparative analyses revealed that the rhizosphere microbiome of M×g was reorganised under heavy metal, hydrocarbon, and mixed contamination, with patterns of change depending on the type of pollutant. In Zn-spiked soil, overall microbial diversity and the abundance of unique taxa decreased, consistent with previous reports [78]. The most pronounced decreases were observed in Bacillus (Firmicutes), Rubrobacter (Actinobacteriota), Vicinamibacter (Acidobacteriota), and the WD2101_soil_group (Planctomycetota). Conversely, Zn contamination stimulated the enrichment of Pseudarthrobacter, Micromonospora, Nocardioides (Actinobacteriota), Sphingomonas, Methylobacterium-Methylorubrum, Aquicella (Alphaproteobacteria), JG30-KF-CM45, C0119 (Chloroflexota), and Bryobacter (Acidobacteriota) (Figure 3). Similar metal-induced shifts in microbial community structure have been documented [22,43,44,46]. For instance, Bourgeois et al. [43] found that M×g cultivation on soils contaminated with persistent organic pollutants and heavy metals (Zn, Pb, Cu, and Cd) promoted the growth of Rhizobiales, Nitrospira, Azospira, and Gemmatimonas, as well as fungi of the phyla Glomeromycota and Mortierella. Pham et al. [44] reported that heavy metal contamination shifted rhizosphere bacterial communities towards opportunistic genera such as Pseudomonas and Stenotrophomonas, though with only minor effects on overall diversity and richness. Other studies have shown that bacterial diversity in the rhizosphere, rhizoplane, and endosphere of M×g remains relatively stable under metal exposure, despite strong uptake by roots [46]. Enrichment of Luteolibacter and Micromonospora in the endosphere under heavy metal stress has been linked to improved plant performance [46]. Our findings align with these observations, particularly the enrichment of Micromonospora in the M×g rhizosphere under Zn contamination (Figure 3). Although resistance mechanisms of Micromonospora to the metalloid TeO32− have been described [79], their bioremediation potential remains largely unexplored.
Several taxa stimulated by Zn in our study are known to play dual roles in remediation and plant growth promotion. These include Bacillus [80], Sphingomonas [35,81], Pseudarthrobacter [82,83], and Micromonospora [84], all of which exhibit metal resistance and contribute to phytostabilisatioin or phytoaccumulation processes.
Soil contaminated with oil sludge exhibited the lowest diversity and the fewest unique taxa. The abundance of Bacillus (Firmicutes), Micromonospora, Pseudarthrobacter, Microlunatus, and Rubrobacter (Actinobacteriota), JG30-KF-CM45, KD4-96, and C0119 (Chloroflexota), and Gemmatimonas (Gemmatimonadota) declined most notably under oil sludge contamination. In contrast, taxa stimulated by oil sludge included Candidatus_Koribacter, uncultured Acidobacteriales, Bryobacter (Acidobacteriota), Mycobacterium (Actinobacteriota), Sphingomonas, KCM-B-112, Parvibaculum, Acidibacter, Immundisolibacter, Mesorhizobium, as well as representatives of Xanthobacteraceae and Oxalobacteraceae (Pseudomonadota) and Candidatus_Udaeobacter (Verrucomicrobiota). Notably, Sphingomonas [85,86] and Mycobacterium [87,88] are established hydrocarbon degraders.
Our results on changes in the rhizosphere bacterial community of M×g are consistent with previous findings. Nebeská et al. [48] reported a decline in Actinobacteriota, a predominant phylum, and an increase in Verrucomicrobia, Bacterioides, and Acidobacteriota in the rhizosphere community of M×g grown in oil-contaminated soil. Hydrocarbon degraders such as Pseudomonas, Shinella, Altererythrobacter, Azospirillum, Mesorhizobium, and Dyella were also enriched in Miscanthus-planted soils compared with unplanted controls [48].
Mixed contamination with Zn and oil sludge supported the highest diversity and the largest number of unique taxa. Besides a reduction in Bacillus, the most prominent feature was the enrichment of Proteobacteria members including Sphingomonas, KCM-B-112, Rhodanobacter, Burkholderia-Caballeronia-Paraburkholderia, Mesorhizobium, Dongia, and Pseudomonas. Comparative analysis of treatments suggested that taxa such as Rhodanobacter and Pseudarthrobacter were primarily stimulated by Zn contamination, whereas oil sludge promoted KCM-B-112, Sphingomonas, and Mycobacterium. Unique taxa in co-contaminated soils = included Burkholderia-Caballeronia-Paraburkholderia, Mesorhizobium, Dongia, and Pseudomonas.
Although only 2–5% of rhizosphere microorganisms directly promote plant growth [89], plants selectively recruit beneficial taxa under stress [90]. Our metagenomic analysis revealed several plant growth–promoting taxa in the M×g rhizosphere, including Bacillus (Bacillaceae) [35,91], Sphingomonas (Sphingomonadaceae) [92], Mesorhizobium (Phyllobacteriaceae) [93], Pseudarthrobacter [82,83], and Micromonospora [84].
In addition to community-level analysis, we isolated strains from various taxa with strong remediation potential. These isolates combined heavy metal resistance and/or hydrocarbon-degrading capacity with plant-growth-promoting activity, making them promising candidates for enhancing M×g performance on contaminated soils. Nonetheless, validation in field conditions is required before practical applications can be confirmed.
Overall, this study characterised the taxonomic composition of the M×g rhizosphere microbiome and its response to soil contamination. Future research should extend beyond taxonomic profiling to include fungal communities and, importantly, functional characterisation using metagenome-assembled genomes (MAGs) and metatranscriptomics. These approaches will enable the linking of microbial taxa to ecologically relevant functions, including plant growth promotion and pollutant resistance or degradation.

5. Conclusions

This comparative study of the M×g rhizosphere microbiome in soils spiked with Zn, oil sludge, or both revealed pollutant-specific reorganisation of the rhizosphere bacterial community. Both contaminants reduced microbial diversity when applied individually, selectively stimulating or inhibiting particular taxa. Oil sludge induced more pronounced taxonomic shifts than Zn and largely determined in the structure of communities under mixed contamination. Specifically, Zn contamination increased the proportions of Actinobacteriota, Pseudomonadota, and Bacteroidota, while decreasing Firmicutes, Planctomycetota, and Acidobacteriota. In contrast, oil sludge promoted Pseudomonadota, Acidobacteriota, and Verrucomicrobiota, but suppressed Actinobacteriota, Bacteroidota, Chloroflexota, Firmicutes, and Gemmatimonadetes. The co-contaminated soil supported the highest diversity and contained the greatest number of unique genera, particularly Gammaproteobacteria, Actinobacteriota, Firmicutes and Bacteroidota.
The rhizosphere community of M×g under co-contamination harboured taxa with hydrocarbon-degrading, heavy metal–tolerant, and plant-growth-promoting traits. Notably, Sphingomonadaceae increased under all contamination scenarios, suggesting their involvement in oil sludge degradation, Zn tolerance, and adaptation to the M×g rhizosphere.
In total, 76 strains were isolated from M×g rhizosphere and screened, of which several exhibited combined metal resistance, hydrocarbon-degrading capacity, and plant growth–promoting activities such as nitrogen fixation, phosphorus mobilisation, siderophore production, and phytohormone synthesis.
Thus, M×g–associated rhizosphere microorganisms represent promising candidates for the development of microbial bioagents to enhance plant growth and improve the efficiency of soil remediation in contaminated environments.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/agronomy15092232/s1. Figure S1: Relationship between the number of detected OTUs and sequencing depth in M×g rhizosphere soil samples; Table S1: Predominant taxonomic composition of rhizosphere bacterial communities of M×g grown in control and differently spiked soils; Table S2: Unique genus-level OTUs in the rhizobiome of M×g grown in control soil; Table S3: Unique genus-level OTUs in the rhizobiome of M×g grown in Zn-spiked soil; Table S4: Unique genus-level OTUs in the rhizobiome of M×g grown in oil sludge-spiked soil; Table S5: Unique genus-level OTUs in the rhizobiome of M×g grown in Zn- & oil sludge-spiked soil.

Author Contributions

Conceptualization, A.M. (Anna Muratova); methodology, I.S.; validation, E.B. and A.M. (Aigerim Mamirova); formal analysis, E.B., I.S., R.B. and A.M. (Anna Muratova); investigation, E.B. and I.S.; resources, A.N.; data curation, E.B., I.S., R.B. and A.M. (Anna Muratova); writing—original draft preparation, A.M. (Aigerim Mamirova) and A.M. (Anna Muratova); writing—review and editing, A.N., E.B., I.S., A.M. (Aigerim Mamirova), R.B. and A.M. (Anna Muratova); supervision, A.N.; project administration, A.N.; funding acquisition, A.N. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Committee of Science, the Ministry of Science and Higher Education, the Republic of Kazakhstan, grant number AP23487419.

Data Availability Statement

The original contributions presented in the study are included in the article, further inquiries can be directed to the corresponding author/s.

Acknowledgments

We are very grateful to Sergey Golubev (IBPPM RAS) for his help in depositing raw reds in the NCBI Sequence Read Archive.

Conflicts of Interest

The authors declare no conflicts of interest.

Abbreviations

The following abbreviations are used in this manuscript:
ABTsAlcohol-benzene tars
DWDry weight
DNADeoxyribonucleic acid
IAAIndole-acetic acid
LBLuria–Bertani medium
LSDLeast significant difference
M×gMiscanthus × giganteus
MAGsMetagenome-assembled genomes
MBCAMono- and bicyclic aromatic hydrocarbons
MPCMaximum permissible concentration
NCBINational Centre for Biotechnology Information
nMDSNonmetric multidimensional scaling
OTUOperational taxonomic unit
PAHsPolycyclic aromatic hydrocarbons
PCAPrincipal component analysis
PCRPolymerase chain reaction
rRNARibosomal ribonucleic acids

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Figure 1. NMDS for M×g rhizosphere bacterial communities: ●—control soil; ▲—Zn-spiked soil; ■—oil sludge-spiked soil; ♦—Zn- & oil sludge-spiked soil.
Figure 1. NMDS for M×g rhizosphere bacterial communities: ●—control soil; ▲—Zn-spiked soil; ■—oil sludge-spiked soil; ♦—Zn- & oil sludge-spiked soil.
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Figure 2. Relative abundance of bacterial communities at the phylum level in the rhizosphere of M×g grown in control and contaminated soils. Note: OS—oil sludge.
Figure 2. Relative abundance of bacterial communities at the phylum level in the rhizosphere of M×g grown in control and contaminated soils. Note: OS—oil sludge.
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Figure 3. Effect of Zn and oil sludge on the relative abundance of predominant genus-level OTUs in the rhizosphere of M×g. Values significantly different from the control are indicated in red (higher) or blue (lower) at p ≤ 0.05.
Figure 3. Effect of Zn and oil sludge on the relative abundance of predominant genus-level OTUs in the rhizosphere of M×g. Values significantly different from the control are indicated in red (higher) or blue (lower) at p ≤ 0.05.
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Figure 4. Venn diagram of common and unique (in parentheses) OTUs at the genus level in the rhizosphere bacterial communities of M×g grown in control and contaminated soils.
Figure 4. Venn diagram of common and unique (in parentheses) OTUs at the genus level in the rhizosphere bacterial communities of M×g grown in control and contaminated soils.
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Table 1. Soil agrochemical profiles.
Table 1. Soil agrochemical profiles.
ParameterUnitControl SoilZn-Spiked SoilZn- & Oil Sludge-Spiked SoilOil Sludge-Spiked Soil
pH−5.645.375.545.53
NO3-Nmg kg−137.8 ± 6.2918.6 ± 0.7117.0 ± 3.2420.6 ± 0.20
NH4-Nmg kg−131.7 ± 1.5539.6 ± 2.1431.7 ± 1.8731.8 ± 6.60
P2O5mg kg−163.8 ± 0.9851.0 ± 6.5353.2 ± 5.5558.3 ± 4.57
Table 2. The α-diversity indices for M×g rhizospheric microbial communities.
Table 2. The α-diversity indices for M×g rhizospheric microbial communities.
TreatmentObserved FeaturesChao1Shannon IndexSimpson IndexFaith PD
Control soil1054 ± 1481062 ± 1478.74 ± 0.180.994 ± 0.00182.5 ± 8.96
Zn-spiked soil709 ± 130 *718 ± 135 *8.46 ± 0.390.995 ± 0.00264.3 ± 9.64
Oil sludge-spiked soil575 ± 202 *582 ± 208 *8.26 ± 0.430.995 ± 0.00149.3 ± 14.8 *
Zn- & oil sludge-spiked soil814 ± 207822 ± 2118.77 ± 0.350.997 ± 0.001 *71.9 ± 15.8
Notes: PD—phylogenetic diversity; *—significantly different compared control at p < 0.05.
Table 3. Microbial isolates with potential applications in bioremediation.
Table 3. Microbial isolates with potential applications in bioremediation.
IsolateN2 FixationP MobilizationSiderophore ProductionIAA SynthesisResistance, mMHC Degradation
Zn2+Pb2+
Chitinophaga sp. Zn19+–++4.0––
Mycolicibacterium sp. Pb113+–++–≥5.0+ (Phe)
Stenotrophomonas sp. Zn210–––±≥5.0–+ (alk)
Mesorhizobium sp. Pb210+++±+–2.5+ (Phe)
Mycolicibacterium sp. Pb216+–++–3.0+ (Phe)
Microbacterium sp. Pb24++–+–3.0–
Notes: HC—hydrocarbons; Phe—degradation of phenanthrene in Kiyohara’s test; alk—degradation of paraffines on Bushnell–Haas’s medium.
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Nurzhanova, A.; Boulygina, E.; Sungurtseva, I.; Mamirova, A.; Berzhanova, R.; Muratova, A. Miscanthus × giganteus Rhizobacterial Community Responses to Zn and Oil Sludge Co-Contamination. Agronomy 2025, 15, 2232. https://doi.org/10.3390/agronomy15092232

AMA Style

Nurzhanova A, Boulygina E, Sungurtseva I, Mamirova A, Berzhanova R, Muratova A. Miscanthus × giganteus Rhizobacterial Community Responses to Zn and Oil Sludge Co-Contamination. Agronomy. 2025; 15(9):2232. https://doi.org/10.3390/agronomy15092232

Chicago/Turabian Style

Nurzhanova, Asil, Eugenia Boulygina, Irina Sungurtseva, Aigerim Mamirova, Ramza Berzhanova, and Anna Muratova. 2025. "Miscanthus × giganteus Rhizobacterial Community Responses to Zn and Oil Sludge Co-Contamination" Agronomy 15, no. 9: 2232. https://doi.org/10.3390/agronomy15092232

APA Style

Nurzhanova, A., Boulygina, E., Sungurtseva, I., Mamirova, A., Berzhanova, R., & Muratova, A. (2025). Miscanthus × giganteus Rhizobacterial Community Responses to Zn and Oil Sludge Co-Contamination. Agronomy, 15(9), 2232. https://doi.org/10.3390/agronomy15092232

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