1. Introduction
Karst landforms cover over 12% of the Earth’s total land area, with more than 900,000 square kilometers in China [1]. Karst rocky desertified environments are typical ecosystems [2] with high sensitivity, low environmental capacity, weak resistance to disturbance, poor stability, vulnerability, poor recovery capability, soil rich in calcium and magnesium and poor nutrients [3,4]. There are two substrate types with highly specific physico-chemical properties in the soil of such areas. The limestone substrate is alkaline, rich in calcium and magnesium, extremely poor in essential plant nutrients and prone to fissures upon corrosion of the soluble rock [5,6]. In particular, human activities aggravate environmental problems such as rocky desertification, land resource loss and ecosystem degradation in karst areas, making karst areas vulnerable to ecological environment [6,7], seriously affecting plant growth and limiting vegetation productivity [8].
The introduction of leguminous trees capable of forming symbiotic relationships with nodulating nitrogen (N)-fixing bacteria and arbuscular mycorrhizal fungi (AMF) represents an effective strategy for accelerating soil reclamation and initiating natural succession [9]. These symbiotic associations endow leguminous species with an enhanced ability to grow rapidly in nutrient-poor substrates and to endure the challenging conditions prevalent in degraded soils. Furthermore, most leguminous tree species can establish symbioses with AMF, which are known to interact synergistically with rhizobia, thereby improving plant performance [10,11,12]. The nitrogen fixation capability of these trees also contributes to increased soil bacterial biomass, recovery of soil quality and adaptation to Karst rocky desertification environments, thus offering significant potential for expediting Karst rocky reclamation [13,14]. Historically, emphasis has been placed on the nitrogen fixation and AMF associations of nitrogen-fixing species, often overlooking the role of bacteria in the recovery process. However, bacterial communities may be as crucial as AMF for Karst restoration. Karst rocky soils provide an ideal system for examining local adaptation and the relative contributions of plant identity and soil chemistry to bacterial community dynamics.
In the course of long-term synergistic evolution, plants have developed close relationships with key relevant bacteria. These bacteria not only promote plant growth, but they enhance nutrient use, tolerance to biotic and abiotic stresses and plant productivity [15,16]. For instance, the application of some probiotic bacteria isolated from mycorrhizal or rhizospheric soil to the roots of some forest trees significantly improved the growing environment, increased the resistance of seedlings to biotic and abiotic stresses and increased the growth rate of seedlings [17]. Bacteria in rhizospheric soil improve soil ecosystem function and increase plant productivity by increasing the plant absorbed nutrients [18]. There are many factors affecting the bacterial community. There are differences in the structure of the bacterial community in different parts of the same plant, different development stages of the same part and different habitats. For example, roots (apical, middle and rear sections), leaves, flowers, seeds and even nodulated roots may differ from non-nodulated roots. The root tip is the most active part of the root system, releasing the most exudates and significantly influencing the bacterial community structure, thereby enhancing plant growth potential [19].
Here, we studied bacteria in the rhizosphere and root group of a nitrogen (N)-fixing tree species, Dalbergia odorifera. The great N2 fixation efficiency [20] and soil quality improvement of D. odorifera have been reported in our previous studies [21,22,23]. The structure and diversity of the bacterial community in three different parts of the root system (endosphere, rhizosphere and non-rhizospheric parts) in the gravel (similar to karst soil) and the soil substrates were explored using 16S rRNA high-throughput sequencing technology. Subsequently, the physicochemical properties of the substrates were analyzed to investigate the relationship between root-associated microorganisms, plant characteristics and the physicochemical properties of the substrates. The aim of this study was to demonstrate that N-fixing species can adapt to karst soils by altering the endosphere, rhizosphere and non-rhizosphere bacterial community structure, which provides a theoretical basis for the use of D. odorifera for ecosystem recovery and vegetation restoration. We hypothesize that N-fixing tree species a) stimulate bacterial growth in sandy soils; b) have the potential to adapt to karst soils by improving bacterial communities.
2. Materials and Methods
2.1. Sample Sources
We selected live D. odorifera seedlings (34.0 ± 2.0 cm in height and 0.6 ± 0.2 cm in ground diameter) from Bagui nursery, a certified nursery in Guangxi Zhuang Autonomous Region, China. Soil (dark brown limestone loam) and gravel (limestone gravel, ~0.51 cm particle size) were collected from three sites in Mashan County’s rocky desertification area, Guangxi [5,24], to simulate limestone lava fissures. A randomized sampling method ensured representativeness. The soil was homogenized and stored at 4 °C to preserve its properties. Detailed physicochemical properties are in Table 1.
2.2. Experimental Design
The experiment site was the teaching practice base of the School of Forestry, Guangxi University (108°29′ E, 22°86′ N), which has a typical subtropical monsoon climate, an annual solar radiation of 90 kcal-cm−2 to 130 kcal-cm−2, annual sunshine hours of 1169 h to 2219 h, average annual temperature of approximately 21.6 °C and extreme maximum and minimum temperatures of 40.4 °C and −2.4 °C, respectively. The test period of this study was from September 2020 to December 2020, during which the average temperature was 18.6 °C.
The control variate method is used to ensure that all factors remain the same except for the single variable of the culture substrate. The seedlings were placed uniformly in a well-lit outdoor seedbed with an automatic irrigation system where light, air and heat were all consistent. The automatic irrigation system ensured consistent moisture levels across all treatments. Rhizospheric, non-rhizospheric and endospheric bacterial abundance and community composition were analyzed globally and locally under different substrates based on whole-root and two-chambered split-root systems, i.e., whole gravel (R), whole-soil (S), split-soil (DS) and split-gravel (DR) substrates (Figure 1). Planting containers were seedling pots (15 cm in width and 35 cm in height) and split-root pots. The main roots were cut off prior to seedling planting. After rinsing the roots three times with sterile distilled water, the roots were planted into prepared seedling pots and split-root pots, with one seeding in a single container. There were fifteen replicates for each treatment, with a total of fifteen seedlings in this study. Moreover, the seedlings were repositioned every 15 days to reduce the effect of uneven environmental factors.
2.3. Sampling Methods
After uprooting the D. odorifera seedlings by the root shaking method [25], soil samples and gravel in contact with the root system were collected. For non-rhizospheric samples, the substrate was collected directly away from the root system (Figure 1). The roots were washed with running tap water to remove adhering soil. After rinsing thrice with distilled water, the D. odorifera roots were cut into small sections for surface disinfection. Samples from each tissue were immersed in 70% ethanol for 4 min and washed with fresh 2% sodium hypochlorite solution for 1–2 min. Subsequently, samples were immersed in 70% ethanol for 1 min and washed thrice with sterile distilled water [26]. Samples from the rhizosphere were classified into groups as follows: Re, gravel-endosphere; Rr, gravel-rhizosphere; Rn, gravel-non-rhizosphere; SE, soil-endosphere; Sr: soil-rhizosphere; Sn: soil-non-rhizosphere (D, division; R, root; S, soil; e, endosphere; r, rhizosphere; n, non-rhizosphere). The above operation was repeated five times for each group of samples. Due to the inability to tightly adhere soil to roots, the shaking root method was unsuitable. Rhizosphere soil, defined as soil within 4 mm of the root surface [27], was collected, transported to the lab and stored at −80 °C for DNA extraction.
2.4. Total DNA Extraction and Polymerase Chain Reaction Amplification
Genomic DNA was extracted from the samples using the cetyl trimethyl ammonium bromide method [28]. Then, the purity and concentration of the extracted DNA were checked by 1.5% agarose gel electrophoresis and Nanodrop 2000. If the concentration of DNA was too high, the samples were diluted to 1 ng/µL using sterile water. The polymerase chain reaction (PCR) system setting (30 μL) for the specific amplification of root-associated bacteria of D. odorifera was as follows: 10 μL of DNA template, 3 μL each of primers 505F (5′-GTGYCAGCMGCCGCGGTAA-3′) and 806R (5′-GACTACHVGGGTWTCTAAT), 15 μL of 2× Phusion Master Mix and 2 μL of ddH2O. The PCR reaction conditions were as follows: pre-denaturation at 98 °C for 1 min, denaturation at 98 °C for 10 s, annealing at 50 °C for 30 s, extension at 72 °C for 30 s (30 cycles of these four procedures), extension at 72 °C for 5 min and storage at 4 °C. The PCR products were checked for quality by electrophoresis using a 2% concentration agarose gel; those that passed the check were purified by magnetic beads and quantified by enzyme labeling. The PCR products were mixed in equal amounts according to the concentration and then detected by electrophoresis using a 2% agarose gel after thorough mixing. Subsequently, the products were recovered using the GeneJET Gel Recovery Kit (Thermo Scientific, Waltham, MA, USA). Libraries were constructed using an Illumina TruSeq DNA PCR-Free Library Preparation Kit and the constructed libraries were subjected to Qubit quantification and library testing. After passing the testing, the libraries were sequenced using an Illumina NovaSeq 6000 system (San Diego, CA, USA).
By detecting the 16S rRNA V3–V4 region, a total of 6,105,642 raw tags were obtained in this study. Among them, 5,901,731 effective tags were used for subsequent analysis after removing chimeras, with a total of 1,588,455,903 bp of bases and an average length of 414.8 bp per effective tag. The base quality values, Q30 and Q20, were both greater than 92%, indicating reliable data (Table 2).
2.5. Data Statistics and Analysis
The sequencing reads from each sample were spliced using FLASH software and the resulting spliced sequences were defined as raw tags. Further filtering using Qiime 1.9.1 software yielded clean tags. Then, removing chimeras yielded the final effective tags. Each unique sequence in each sample was clustered at 97% into operational taxonomic units (OTUs). After removing the low-abundance OTUs using UCHIME, phylogenetic trees were constructed and, finally, redundancy analysis (RDA) was carried out using Canoco 5.0 to analyze the relationship between bacterial diversity and the physicochemical properties of the substrate.
3. Results
3.1. Bacterial Community Structure and Abundance
The bacterial community structure and diversity of these samples were analyzed at different taxonomic levels according to the OTU annotation results. A total of 79 phyla, 161 classes, 338 orders, 488 families and 868 genera were identified in all samples (Table S1). At the phylum level, Proteobacteria was the dominant phylum in all samples, with the highest percentage of 68.54% in Re. In addition, Cyanobacteria was the second most dominant phylum in Se, DSe and DRe, while Bacteroidota, Actinobacteriota and Acidobacteriota were also present in large numbers in each sample (Figure 2). At the genus level, Herbaspirillum and unidentified Chloroplast comprised the dominant genera in the endosphere of D. odorifera with ≥30% relative abundance of unidentified Chloroplast in the Se, DSe and DRe groups. Similarly, Herbaspirillum was dominant in Rr and DRr, with abundance levels of 20.50% and 10.39%, respectively. However, their abundance levels were low in the Sr and DSr groups (Figure 2). The phylogenetic relationships of the dominant bacteria at the genus level are shown in Figure 3. The dominant phyla observed include Proteobacteria, Cyanobacteria, Bacteroidota, Acidobacteriota, Actinobacteria and Chloroflexi. Proteobacteria show the highest abundance in most groups; other phyla, such as Firmicutes and Verrucomicrobiota, are also present but in lower proportions (Figure 3).
3.2. Alpha Diversity Analysis
The OTUs were classified using a 97% similarity level as the threshold. An amount of 402, 607, 465.8 and 346.2 OTUs were found for the Se, Re, DSe and DRe groups, respectively; 2032.2, 2491.2, 2445.8 and 2433.6 OTUs for Sr, DSr, Sn and DSn, respectively; and 1092.8, 909.6, 675.4 and 1469.2 OTUs for Rr, DRr, Rn and DRn, respectively. The Shannon, Simpson, Chao1, ACE, Goods coverage and PD whole tree indices were used to compare the bacterial diversity of the different samples (Table 3). Sr, DSr, Sn and DSn had the highest Shannon and Simpson indices. In contrast, Se, Re, DSe and DRe had the lowest Shannon and Simpson indices. The Chao1, ACE, Goods coverage and PD whole tree indices showed the same trend, indicating that the bacterial diversity within the endosphere of D. odorifera is low, whereas that in the rhizosphere and non-rhizospheric parts are high.
3.3. Beta Diversity Analysis
To compare the composition and similarity of root-associated bacteria in different samples of D. odorifera, principal coordinates analysis was performed on the bacterial community of each sample based on the Unweighted Unifrac distance. The results showed that the bacteria within the endosphere (Se, Re, DRe and DSe) were more similar and all clustered in the second quadrant. However, all of the Sr, Sn, DSr and DSn in the soil samples clustered in the first quadrant, indicating a high degree of similarity among them (Figure 4). The UPGMA clustering tree also showed the same results. The samples could be divided into three branching clusters based on the distance index between samples, with the Se, Re, DRe and DSe groups clustered in one branch, indicating some similarity among them. The rhizospheric and non-rhizospheric bacteria (Sr, DSr, Sn and DSn) in the soil were clustered in one branch, whereas those in the gravel (Rr, DRr, Rn and DRn) were clustered in another branch (Figure 5). These results suggest that the type of substrate strongly influenced rhizospheric and non-rhizospheric bacterial communities and their endophytes are largely determined by plant species.
3.4. Relationship Between Root-Associated Bacteria and Physicochemical Properties of Substrates
To investigate the relationship between physicochemical properties and the relative abundance of the dominant bacteria, an RDA of the dominant bacteria and substrate physicochemical properties in the root system of D. odorifera was conducted, as shown in Figure 6. At the phylum level, the lateral axis was positively correlated with Proteobacteria and Cyanobacteria and negatively correlated with Bacteroidota, Firmicutes, Actinobacteriota, Chloroflexi, Desulfobacterota and WPS-2, explaining 66.42% of the information (Figure 6a). The vertical axis was positively correlated with Acidobacteriota, explaining 17.22% of the data. According to the RDA plot, the dominant bacterium Proteobacteria showed a significant positive correlation with total carbon (TC) and pH value and a negative correlation with available phosphorus (AP), moisture content (MC), organic matter (OM), available kalium (AK) and total nitrogen (TN). Notably, the Monte Carlo test showed that MC, pH value and AP were significantly correlated with bacterial community composition at the phylum level (p < 0.05) (Table 4).
At the genus level, the lateral axis was positively correlated with Herbaspirillum, Chloroplast, Rhizobium, Pseudomonas and Bradyrhizobium and negatively correlated with Aquabacterium and Sphingomonas, explaining 68.48% of the data (Figure 6b). The vertical axis showed a positive correlation with Acinetobacter and a negative correlation with Chitinophaga, explaining 8.56% of the data. In addition, Herbaspirillum and MC elements were in the same direction at the same time, indicating a significant positive correlation between MC elements and Herbaspirillum. Similarly, Acinetobacter was in the same direction as pH value, indicating that pH value had a positive correlation with Acinetobacter; Acinetobacter was in the opposite direction to AK, MC, OM, TN and AP, indicating a negative correlation between them. In contrast, Bradyrhizobium, Chitinophaga and Pseudomonas were in the same direction as AK, MC, OM, TN and AP, suggesting a positive correlation between them. Notably, the Monte Carlo test showed that pH value and MC were significantly correlated with bacterial community composition at the genus level (p < 0.05) (Table 4).
4. Discussion
Consistent with other studies, rhizosphere bacterial diversity is much lower than that of the inner sphere [29]. Furthermore, endospheric bacterial diversity was not influenced by different substrates, whereas rhizospheric and non-rhizospheric bacterial diversity was more influenced by the type of substrates. Interestingly, the significantly lower bacterial abundance in the rhizosphere and non-rhizosphere soils (S and DS treatments) of N-fixing plants compared to stony substrates may be attributed to multiple environmental and ecological factors. First, stony substrates (e.g., gravel or rocky matrices) often exhibit higher porosity and better aeration, which enhance oxygen diffusion and create favorable microhabitats for aerobic bacteria, including N-fixing species [30]. In contrast, soil matrices, particularly in the rhizosphere, may experience localized hypoxia due to root respiration and microbial activity, limiting bacterial proliferation [30]. Second, nutrient competition in the rhizosphere is intensified by root exudates and plant–microbe interactions. While root exudates provide carbon sources, they also stimulate competitive exclusion among microbial communities, potentially suppressing certain bacterial populations [31]. In stony substrates, reduced competition for resources and lower organic carbon availability might favor specialized bacterial taxa adapted to oligotrophic conditions. Third, physicochemical properties of stony substrates, such as higher thermal stability and reduced water retention, could selectively enrich bacterial groups resistant to desiccation and temperature fluctuations [32]. Soil environments, by contrast, undergo more dynamic moisture and temperature changes, which may stress bacterial communities. However, the rhizospheric zone is a highly dynamic and complex area of chemical, physical and biological interactions, which is constantly receiving secondary metabolites [33]. The results need to be verified from root exudations and other aspects in the future.
Root-associated bacteria in plants play a crucial role in plant growth by fixing nitrogen and solubilizing phosphorus, potassium and plant-growth-regulating hormones. Among these, Proteobacteria represent one of the most diverse and rapidly metabolizing phyla of bacteria, contributing to the soil ecological stability by facilitating the nitrogen cycle and efficiently utilizing various substances secreted from the plant rhizosphere [34]. This is consistent with the results of previous studies that indicate the abundance of Proteobacteria in karst areas is higher than other bacteria [35]. The high relative abundance of Proteobacteria in all samples across this study suggests their adaptability to the diverse environments associated with D. odorifera roots. This adaptability is particularly significant in the context of N and P cycling, as Proteobacteria are known to play a crucial role in the mineralization of organic N and P compounds, thereby enhancing nutrient availability for plant uptake. The interaction between Proteobacteria and D. odorifera roots is further supported by their ability to metabolize root exudates, which are rich in carbon compounds that fuel microbial activity in the rhizosphere [36]. Moreover, the co-occurrence of Proteobacteria with other N-fixing bacteria, such as Rhizobium and Bradyrhizobium, can lead to increased nitrogen fixation rates, which are critical for the growth of leguminous plants like D. odorifera [37]. For instance, the co-occurrence of Proteobacteria with other N-fixing bacteria, such as Rhizobium and Bradyrhizobium, can lead to increased nitrogen fixation rates, which are critical for the growth of leguminous plants like D. odorifera [37].
Acidobacteria also exhibited broad adaptability and functional diversity [38]. The observed higher relative abundance of Acidobacteria in rhizospheric and non-rhizospheric samples compared to endospheric samples suggests that Acidobacteria present the endosphere may originate from the substrate. Additionally, Actinobacteria, which are Gram-positive bacteria, play a significant role in the degradation of cellulose and chitin, thereby contributing to soil nutrient supply. They also possess a normal metabolic rate and the ability to repair DNA under extreme conditions [38]. The relative abundance of Actinobacteria was found to be significantly higher under gravel conditions, because it can produce various hydrolytic enzymes and secondary metabolites, which help them compete for resources in low-nutrient environments [39]. Compared to soils, the gravel substrate exhibited a more pronounced nitrogen deficiency, a nutrient-limiting condition that may selectively promote the growth of Actinobacteria, as they typically possess efficient nitrogen acquisition and utilization capabilities, such as nitrogen fixation or the decomposition of complex organic nitrogen compounds. This finding aligns with our research results (Figure 5) and is consistent with previous studies. Firmicutes also emerged as one of the dominant bacterial taxa across all sample groups, exhibiting the highest abundance in both rhizospheric and non-rhizospheric soil samples. Previous research has indicated a correlation between Firmicutes content and carbon levels [40], which may have influenced the relative abundance of Firmicutes in the rhizospheric bacteria of D. odorifera. Additionally, Chloroflexi, a Gram-negative bacterium capable of utilizing light energy for photosynthesis to sustain autotrophic growth [41], was identified as another dominant bacterial community in both rhizospheric and non-rhizospheric regions across different substrates. This suggests that Chloroflexi plays a significant role in the ecological cycling of soil.
At the genus level, Herbaspirillum was identified as the predominant bacterium within the endosphere. As a class of N-fixing bacteria, Herbaspirillum is implicated in the N fixation process in D. odorifera. It does not exert any deleterious effects on the host plant, but rather enhances crop yield and nitrogen content and promotes nutrient uptake by plants through complex molecular exchanges with them [42,43,44,45]. In addition, Herbaspirillum was also the dominant bacterium in the rhizosphere of plants cultivated in gravels, exhibiting significantly higher abundance compared to non-rhizospheric samples in gravels and endospheric samples grown in soils; this may be related to the N fixation of D. odorifera. Our previous study found that the nitrogen fixing efficiency of D. odorifera was negatively correlated with soil nitrogen nutrition, which was strongly confirmed by the high N content of soil in this study [20]. Further, Sphingomonas is a Gram-negative bacterium capable of decomposing organic compounds in the substrate was predominantly found in the rhizosphere and non-rhizospheric regions of the samples. Similarly, Acinetobacter was one of the dominant bacteria in the rhizosphere and non-rhizospheric regions of D. odorifera. Acinetobacter is known to inhibit the activity of pathogenic bacteria and has been reported as a potential biocontrol agent. The organism studied promotes plant growth by producing growth hormones, increasing phosphate levels, synthesizing iron chelators and producing ammonia [46]. Another significant Gram-negative bacterium, Pseudomonas, predominantly found in soil, water and various plant tissues, exhibits a robust capacity to decompose organic matter and utilize a diverse array of organic compounds as energy sources [47].
The composition of root-associated bacterial communities in plants is subject to influence from soil-dwelling bacteria [48] as well as various environmental factors. Factors such as soil texture, the plant growth environment, nature of the internal microenvironment of plants, gene levels and human production activities all affect the community structure of endophytic fungi [49,50]. The results from RDA showed that the structure of the root-associated bacterial community is primarily influenced by soil C, pH and available phosphorus levels. We also found Proteobacteria showed a significant positive correlation with both soil pH and soil C content, but negatively correlated with available P. Although previous research identified a significant positive correlation between Proteobacteria and total soil potassium in Eucalyptus forest ecosystems [49,51], they are also affected by other soil properties because they are highly competitive in utilizing available nutrients resources in nutrient-poor soils [52]. These results show that soil nutrients significantly shape the microbial habitat and resource availability. Specifically, soil organic carbon serves as a critical carbon source for microorganisms, fueling their metabolic activities and promoting microbial growth and activity. Soil pH significantly influences microbial community composition by affecting enzyme activity and nutrient availability. Additionally, available nutrients, such as nitrogen and phosphorus, shape microbial dynamics by determining resource allocation and metabolic pathways, leading to shifts in community structure and function [53]. Together, these factors create a complex interplay that governs microbial diversity and ecosystem processes. Our results also demonstrate that rhizosphere microbial communities of N-fixing tree species still adhere to these rules in karst soils. When substrate nutrients are low, microorganisms may be stimulated to adapt to this environment, which is directly confirmed by our results that bacterial abundance in S and DS treatments is lower than that in R and DR treatments, respectively.
Chloroflexi, an autotrophic phylum that operates independently of external nutrient sources [54], is prominently recognized as a bioindicator of suboptimal soil conditions due to its frequent occurrence in such environments. This phylum demonstrates pronounced sensitivity to variations in soil pH, exhibiting a significant negative correlation with substrate pH levels. This relationship highlights the pivotal role of pH in modulating the distribution and ecological function of Chloroflexi. In contrast, a reduced abundance of Acidobacteria is often indicative of enhanced soil quality, as this phylum predominantly thrives in nutrient-deficient soils [49]. Meanwhile, Bacteroidetes play an essential role in nutrient cycling and the dynamics of soil organic matter, attributed to their proficiency in decomposing organic matter, thereby enhancing soil fertility and promoting plant growth. It is noteworthy that when soil ammonium nitrogen levels are sufficient to support both plant and microbial growth, the decomposition of organic matter mediated by Bacteroidetes diminishes significantly [49]. This observation suggests a metabolic shift within microbial communities in response to varying nitrogen availability. Our study’s substantial Bacteroidetes abundance highlights the adaptability of N-fixing tree species in nutrient-poor karst ecosystems through microbial community restructuring. These findings underscore the complex relationships among microbial taxa, soil properties and nutrient availability. The results emphasize the necessity of adopting a comprehensive approach to understanding microbial community dynamics for accurate assessment of soil health and ecosystem functionality in fragile karst environments.
5. Conclusions
This study elucidates the profound influence of culture substrates on the root system architecture and rhizospheric bacterial community of D. odorifera through the application of split-root systems. The key findings reveal that bacterial communities in the endosphere, rhizosphere and non-rhizosphere of N-fixing plants exhibit remarkable resilience in nutrient-poor karst environments, with their composition and dynamics strongly correlated with soil nutrient availability, pH and carbon sources. These findings provide robust evidence that plant–microbe interactions play a pivotal role in root development and the assembly of microbial communities. Moreover, the study highlights D. odorifera as a highly adaptable species for ecological restoration in karst regions, with its root-associated bacterial communities demonstrating exceptional adaptability to challenging environmental conditions. Collectively, these insights deepen our understanding of plant–microbe–soil interactions and underscore the potential of D. odorifera in sustainable ecosystem restoration.
Conceptualization, S.Y. and X.Y.; methodology, X.Y. and C.G.; formal analysis, C.G.; resources, S.Y.; writing—original draft preparation, C.G. and X.Y.; writing—review and editing, X.Y.; funding acquisition, S.Y. and X.Y. All authors have read and agreed to the published version of the manuscript.
Data are available from corresponding author upon reasonable request.
The authors declare no conflicts of interest.
Footnotes
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Enlarge this image.Figure 1. Schematic representation of four treatments and three root zones, including endosphere, rhizosphere and non-rhizospheric zones; S: soil; R: rock; D: division.
Enlarge this image.Figure 2. Horizontal histogram of the root-associated bacterial of D. odorifera at the genus level. Sample abbreviations in the figure: Re, grave-endosphere; Rr, gravel-rhizosphere; Rn, gravel-non-rhizosphere; SE, soil-endosphere; Sr: soil-rhizosphere; Sn: soil-non-rhizosphere (D, division; R, root; S, soil; e, endosphere; r, rhizosphere; n, non-rhizosphere).
Enlarge this image.Figure 3. Phylogenetic analysis of the rhizospheric bacteria of D. odorifera (top 100). Sample Abbreviations in the figure: Re, gravel-endosphere; Rr, gravel-rhizosphere; Rn, gravel-non-rhizosphere; SE, soil-endosphere; Sr: soil-rhizosphere; Sn: soil-non-rhizosphere (D, division; R, root; S, soil; e, endosphere; r, rhizosphere; n, non-rhizosphere).
Enlarge this image.Figure 4. Principal Component Analysis (PCA) analysis of root-associated bacteria of D. odorifera. Sample abbreviations in the figure: Re, gravel-endosphere; Rr, gravel-rhizosphere; Rn, gravel-non-rhizosphere; SE, soil-endosphere; Sr: soil-rhizosphere; Sn: soil-non-rhizosphere (D, division; R, root; S, soil; e, endosphere; r, rhizosphere; n, non-rhizosphere).
Enlarge this image.Figure 5. Cluster analysis (phylum level) of root-associated bacteria of D. odorifera. Sample abbreviations in the figure: Re, gravel-endosphere; Rr, gravel-rhizosphere; Rn, gravel-non-rhizosphere; SE, soil-endosphere; Sr: soil-rhizosphere; Sn: soil-non-rhizosphere (D, division; R, root; S, soil; e, endosphere; r, rhizosphere; n, non-rhizosphere).
Enlarge this image.Figure 6. RDA of the rhizospheric bacteria of D. odorifera. (a): phylum level, (b): genus level. O: Soil organic matter; C: Soil carbon; N: Nitrogen; P: Soil available phosphorus; K: Soil available kalium; W: soil water content.
Physico-chemical properties of substrates.
| pH Value | MC | TC | TN | AP | AK | OM | |
|---|---|---|---|---|---|---|---|
| Soil | 6.21 | 33.1 | 11.88 | 1.01 | 3.6 | 62 | 20.48 |
| Gravel | 9.48 | 0.52 | 12 | 0.11 | 0.15 | 6.49 | 0 |
MC: Moisture Content; OM: Organic Matter; TC: Total Carbon; TN: Total Nitrogen; AP: Available Phosphorus; AK: Available Kalium.
Basic information obtained from sequencing.
| Sample Name | Raw PE | Base (nt) | AvgLen (nt) | Q20 | Q30 | GC (%) | Effective (%) |
|---|---|---|---|---|---|---|---|
| Srhi | 99,941.6 | 25,765,576 | 411.6 | 97.99 | 93.62 | 55.81 | 62.68 |
| DSrhi | 99,930.8 | 25,926,718 | 415 | 97.90 | 93.39 | 55.66 | 62.59 |
| Sendospheric | 103,612.4 | 27,651,788 | 414 | 97.82 | 93.13 | 54.38 | 64.53 |
| Rendospheric | 101,881.4 | 26,363,271 | 413.2 | 97.77 | 93.10 | 53.46 | 62.65 |
| Drhi-endospheric | 102,400.6 | 26,632,252 | 412.6 | 97.79 | 93.12 | 54.39 | 62.96 |
| Rnon-rhi | 104,041.8 | 27,186,515 | 415.4 | 97.79 | 93.17 | 55.29 | 62.93 |
| Snon-rhi | 101,313.2 | 26,286,594 | 413.6 | 97.89 | 93.38 | 55.77 | 62.74 |
| DSendospheric | 104,189.6 | 26,659,058 | 413.2 | 97.82 | 93.18 | 54.18 | 61.93 |
| Rrhi | 103,576.4 | 26,486,781 | 417.6 | 97.72 | 92.91 | 54.62 | 61.27 |
| DSnon-rhi | 100,861.8 | 25,770,447 | 417.2 | 97.81 | 93.15 | 55.22 | 61.31 |
| DRrhi | 100,144.2 | 26,994,511 | 417.8 | 97.77 | 93.04 | 54.59 | 64.96 |
| DRnon-rhi | 99,234.5 | 25,967,669 | 416.25 | 97.76 | 93.08 | 55.04 | 62.83 |
| Total | 610,5642 | 1,588,455,903 | 4977.45 | / | / | / | 62.78 |
Alpha diversity assay.
| Shannon | Simpson | Chao 1 | Ace | Goods Coverage | PD Whole Tree | |
|---|---|---|---|---|---|---|
| Se | 2.888 | 0.742 | 471.154 | 498.328 | 0.998 | 52.454 |
| Re | 4.436 | 0.821 | 665.362 | 673.542 | 0.998 | 77.348 |
Significance test for correlation between physico-chemical properties of substrates.
| Environmental | Phylum Level | Genus Level | ||||
|---|---|---|---|---|---|---|
| Rate of | F | p | Rate of | F | p | |
| pH | 21.9 | 2.3 | 0.136 | 3.0 | 0.2 | 0.020 |
| MC | 22.2 | 2.3 | 0.138 | 3.0 | 0.2 | 0.022 |
| TC | 41.2 | 5.4 | 0.014 | 15.4 | 1.4 | 0.186 |
| TN | 16.1 | 1.6 | 0.228 | 9.0 | 0.8 | 0.054 |
| AP | 22.6 | 2.4 | 0.12 | 2.8 | 0.2 | 0.190 |
| AK | 18.1 | 1.8 | 0.176 | 6.1 | 0.5 | 0.160 |
| OM | 20.3 | 2.1 | 0.132 | 3.9 | 0.3 | 0.198 |
MC: Moisture Content; OM: Organic Matter; TC: Total Carbon; TN: Total Nitrogen; AP: Available Phosphorus; AK: Available Kalium.
Supplementary Materials
The following supporting information can be downloaded at:
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; Guo, Chunhui 2 ; Yao, Xianyu 1 1 Guangxi Key Laboratory of Forest Ecology and Conservation, Guangxi Colleges and Universities Key Laboratory for Cultivation and Utilization of Subtropical Forest Plantation, and State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, College of Forestry, Guangxi University, Daxuedonglu 100, Nanning 530004, China;
2 Ecological Environment Monitoring and Scientific Research Center, Yellow River Basin Ecology and Environment Administration, Ministry of Ecology and Environment, Zhengzhou 450000, China;