Analysis of Microbial Community Structure Around Roots of Stipa Grandis

doi:10.21203/rs.3.rs-500898/v1

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Analysis of Microbial Community Structure Around Roots of Stipa Grandis

https://doi.org/10.21203/rs.3.rs-500898/v1

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The root zone microbial structure is particularly complex for plants with rhizosheaths, which may play an important role in the future agricultural sustainable development. However, one of the important reasons for restricting our study of rhizosheath microbial structure is that there is no definite method for rhizosheath separation. The aim of this study was to explore the isolation methods of rhizosheath and the diversity and functional characteristics of microorganisms around the rhizosphere. In this study, we isolated the rhizosheath of Stipa grandis, a dominant species in desert steppe, and the microorganisms in the roots, root epidermis, rhizosheath, rhizosphere soil were extracted and sequenced by 16s RNA and ITS. The bacterial alpha diversity index was in the order rhizosphere soil > rhizosheath > root epidermis > endophytic, and the fungal alpha diversity index was rhizosphere soil and rhizosheath > root epidermis and endophytic. There were significant differences in bacterial community structure between the root epidermis and endophytic, rhizosheath, rhizosphere soil, and the sum of relative abundance of the dominant bacterial populations Actinobacteria and Proteobacteria was 73.9% in the root epidermis. Different from bacterial community structure, the community structure of root epidermis fungi was similar to endophytic, but significantly different from rhizosheath and rhizosphere soil. We suggest that the root epidermis can act as the interface between the host plant root and the external soil environment. This study will provide theoretical and technical guidance for the isolation of plant rhizosheath and the study of microorganisms in it.

Agricultural Engineering

Environmental Policy

Stipa grandis

Rhizosheath

Rhizosphere

Endophytic

Microorganism

The close interaction between plants and rhizosphere microorganisms has prompted people to regard plants as a superorganism (Cornéet al. 2014). Research on plant root – rhizosphere soil microorganisms aims to identify correlations between plant – soil – microorganisms and reveal their important roles in the ecosystem. However, the relationship between the root and the soil around the root of some plants is more complex, and these plants are plants with rhizosheaths. For example, barley (George et al. 2014), wheat (Delhaize et al. 2015), corn (Duell and Peacock 1985), rushes (Shane et al. 2009, 2011), etc. even some scholars have proposed that there is a rhizosheath on the fine roots of some leguminous plants (Sprent 1975; Unno 2005). However, McCully (1999) suggest that this needs further research and confirmation. It has been more than 100 years (Volkens 1887) since the rhizosheath was first described, initiating study of its structure (Bailey 1997), formation, function and genetic characteristics. However, little is known about it even to this day.

Rhizosheath is particularly obvious in the root structure of Gramineae in arid areas. Price (1911) and young (1995) believe that the rhizosheath plays an important role in increasing the drought resistance of plants, and existing research supports this view (Pate and Dixon, 1996; Shane et al. 2010; Benard et al. 2016). This understanding is of great significance for improving agricultural sustainability in the context of future climate change, limited resources and a growing global population. Some scholars believe that the rhizosheath plants may play an important role in the second green revolution and future agricultural sustainable development (Lynch 2007; Brown et al. 2017).

The research on the structure and function of plant rhizosphere microorganisms has always been a hot topic, although most microorganisms in the environment have not been cultured, with the rapid development of high-throughput sequencing technology, the structure of microorganisms in the environment is gradually becoming known. Previous studies have shown that there are some differences in the composition of plant root microorganisms and rhizosphere and non-rhizosphere soil microorganisms, which directly affect the normal growth of host plants. For example, some high concentrations of molecules released by rhizosphere microorganisms inhibit the elongation of primary roots and promote the formation of lateral roots and root hairs (Zhang et al. 2017). Some rhizosphere bacteria or fungi produce auxin, which directly interferes with auxin signal transduction (Spaepen et al. 2014). Plant rhizosphere microorganisms play an important role in improving crop yield and resistance. Recent reviews have shown that rhizosphere microorganisms and drought resistant crops interact through several different mechanisms to respond to climate change(Zhang et al. 2017). Different plant species or genotypes can select different rhizosphere microbial communities by producing different secretions from their roots. Although there is still uncertainty about global climate change, there are signs that global temperature will continue to rise, and that drought frequency and duration will change in many locations. Plants respond to this stress through their own regulation. A recent review by Vries (2020) found little evidence for a coupling relationship between the drought tolerance mechanism of microorganisms and the functional characteristics of plant drought resistance, highlighting the need for further research. There are few reports on the structure and function of microorganisms in plant rhizosheath. Therefore, it will be a challenge to study the microorganisms around the rhizosheath plants。

York et al. (2016) summarized and defined the generation process and semantics of "rhizosphere", and considered that rhizosheath is a mixture of soil particles adhered by mucus (the secretion of plant roots or microorganisms) (Volkens 1887; George et al. 2014). The epidermal cell layer attached to the rhizosheath was not a part of the rhizosheath, and they called the combination of the epidermal cell layer and the rhizosheath as "rhizoplane" (York et al. 2016). The difficulty of rhizosheath separation is the separation of rhizosheath and root epidermis. And there is no standard method to isolate microorganisms from rhizosheath. In this study, using Stipa grandis as experimental material, we isolated the roots, root epidermis, rhizosheath and rhizosphere soil of Stipa grandis, extracted microbial DNA from roots, root epidermis, rhizosheath and rhizosphere soil, and sequenced 16S RNA and ITS. The purpose of our research is to explore the isolation methods of rhizosheath and root epidermis, and analyze the similarities and differences of microbial communities among the root, root epidermis, rhizosheath and rhizosphere soil of Stipa breviflora, so as to provide new methods and suggestions for the future research of rhizosheath plants.

Overview of the research site

The experiment was carried out in Maodeng pasture (116.03°E-116.50°E, 44.80 ° N-44.82°N), Xilinhot City, Inner Mongolia, China. The area has a temperate arid continental climate, with an elevation of 1055 m. The annual average temperature is 0–1℃, the frost-free period is 90–115 d, and the accumulated temperature greater than or equal to 0℃ is 1800℃.

Test sample collection

Stipa grandis, a typical gramineous plant in desert steppe, is a constructive species. Samples were taken from Maodeng pasture (Stipa grandis desert steppe) in Xilinhot City, Inner Mongolia, in July 2019. We designed 5 replicates and randomly sampled at 5 sites 50 m apart. Three Stipa grandis were collected from each site as one sample of replicates, and the depth of excavation is 20 cm. The mixed samples of plant and soil were put into plastic bags and immediately placed into ice boxes, and brought back to the laboratory for root system, rhizosheath, rhizosphere and non-rhizosphere soil separation. The isolation of root, rhizosheath, rhizosphere and non-rhizosphere soil and the extraction of microbial DNA were carried out at the Sino-Dutch joint laboratory.

Harvest of roots, root epidermis, rhizosheath, rhizosphere

Phosphate buffer is needed before separating plant roots, root epidermis, rhizosheath, rhizosphere soil. Phosphate buffer plays an important role in maintaining microbial activity and does not change the composition of the sampled microorganisms. Phosphoric acid buffer preparation reagent: disodium hydrogen phosphate, sodium dihydrogen phosphate, Silwet L-77. Preparation of phosphate buffer: phosphate buffer 1 Liter (NaH₂PO₄·H₂O: 6.33 g, Na₂HPO₄·7H₂O: 16.5 g), with addition of 200 µl of Silwet L-77 per liter after high temperature and high pressure sterilization.

Wear sterile gloves and sterilize the workspace with 70% EtOH. The forceps and scissors used in the experiment were wiped and disinfected with 70% EtOH. 3 Stipa plants were selected from each sampling site, and remove loose soil by kneading and shaking by hand and by patting the roots on the back of a gloved hand. The shaken soil is the rhizosphere soil sample. The soil that is not shaken off and adheres to the root surface is defined as rhizosheath soil (Fig. 1). Place the root with rhizosphere soil in a 50 mL tube containing 25 mL phosphate buffer. Vortex max 15 s, and filter rhizosphere through a 100 µm nylon mesh cell strainer into an empty 50 mL tube. After centrifuging at 4000×g for 15 minutes, pour out the supernatant. The soil samples left in the 50 ml tube are rhizosheath soil samples. Remove the root hairs on the filtered root samples with tweezers, and carefully separate the filtered roots and outer epidermal cell layer with tweezers. Place them in a 15 ml tube filled with buffer solution, vortex for 30 s, and wash them 3 times. Place the root and outer epidermal cell layer on sterile filter paper and air dry the root and outer epidermal cell layer sample naturally (6 h).

Sequencing process (Novogene company)

(1) Extraction and PCR amplification of microbial genomic DNA

Microbial DNA from rhizosheath, root epidermis and rhizosphere soil samples was extracted with MOBIO DNeasy PowerSoil kit 12888-100. Microbial DNA from the root was extracted with FastDNATM Spin Kit for Soil. Agarose gel electrophoresis was used to detect the purity and concentration of DNA. An appropriate sample of DNA was applied to the centrifuge tube and diluted with sterile water to 1ng/ L. Using diluted genomic DNA as a template, according to the selection of the sequencing region, specific primers with Barcode, Phusion® High-Fidelity PCR Master Mix with GC Buffer from New England Biolabs company, and high fidelity enzyme were used for PCR to ensure the efficiency and accuracy of amplification. The corresponding regions of primers were: 16S V4 region primers (515F and 806R) -- identification of bacterial diversity. ITS1 region primers (ITS5-1737F and ITS2-2043R) --identification of fungal diversity.

(2) Mixing and purification of PCR products

The PCR product was detected by electrophoresis with 2% agarose gel. According to the concentration of PCR product, the samples were mixed equally, and then the PCR products were detected by agarose gel electrophoresis with 2% agarose gel. The gel recovery kit provided by Qiagen company was used to recover the target band.

(3) Library construction and sequencing

Library construction used TruSeq® DNA PCR-Free Sample Preparation Kit (Building Database Kit). The library was quantified by qubit and Q-PCR, and after the library was qualified, it was sequenced by novaseq6000.

Data analysis based on the Analysis Platform of Novogene company

(1) Sequencing data processing

According to the barcode sequence and PCR amplification primer sequence, each sample data was separated from the offline data. After the barcode and primer sequences were cut off, the reads of each sample were spliced with FLASH (V1.2.7, http://ccb.jhu.edu/software/FLASH/) (Magoč et al. 2011), and the splicing sequence was the original tags data (Raw Tags). The raw tags obtained by splicing need to be filtered (Bokulich et al. 2013) strictly to obtain high-quality tags data (Clean Tags). Following Qiime's tags quality control process (V1.9.1, http://qiime.org/scripts/split_libraries_fastq.html) (Caporaso et al. 2010), the effective data were obtained (Effective Tags).

(2) OTU clustering and species annotation

Uparse software (Uparse v7.0.1001, http://www.drive5.com/uparse/) (Haas et al. 2011) was used to cluster all effective tags of all samples. By default, the sequences were clustered into OTUs (Operational Taxonomic Units) with 97% consistency. At the same time, the representative sequences of OTUs were selected. According to the algorithm principle, the sequence with the highest frequency in OTUs was selected as the representative sequence of OTUs. The OTUs sequences were annotated and analyzed using the Mothur method and the SSUrRNA database (Wang et al. 2007) of silva132 (http://www.arb-silva.de/) (Edgar et al. 2013) (the threshold value was set at 0.8 ~ 1). The taxonomic information was obtained and the community composition of each sample was counted at the phylum level. The fast multi sequence alignment was carried out using MUSCLE (Quast et al. 2013) (Version 3.8.31, http://www.drive5.com/muscle/) software, and the phylogenetic relationships of all OTUs representative sequences were obtained. Finally, the data for each sample was homogenized, and the least amount of data in the sample was taken as the standard. The subsequent alpha diversity analysis and beta diversity analysis were based on the homogenized data.

Alpha diversity analysis: Qiime software (version 1.9.1) was used to calculate observed OTUs, Chao1, Shannon, Simpson, ACE and goods coverage index, and the dilution curve was drawn using R software (version 2.15.3), and alpha diversity index was analyzed using R software.

Beta diversity analysis: The UniFrac distance was calculated with Qiime software (version 1.9.1), and the UPGMA sample clustering tree was constructed. NMDS were drawn using R software (version 2.15.3). Vegan software package of R software was used for NMDS analysis. AMOVA analysis used the amova function in Mothur software.

Distribution characteristics of bacterial and fungal communities

Bacterial OUTs (Fig. 2, a): A total of 8760 OTUs were generated by OTUs clustering at 97% similarity level. Among them, 3787 OTUs were common in roots, root epidermis, rhizosheath, and rhizosphere soil. There were 188, 347, 576 and 711 unique OTUs in roots, root epidermis, rhizosheath, and rhizosphere soil, respectively, accounting for 3.68%, 5.98%, 8.48% and 10.06% of their respective total number of OTUs. Fungal OUTs (Fig. 2, b): A total of 1423 OTUs were generated by OTUs clustering at 97% similarity level. Among them, 318 OTUs were common in roots, root epidermis, rhizosheath soil and rhizosphere soil. The number of unique OTUs in roots, root epidermis, rhizosheath soil and rhizosphere soil were 11, 19, 164 and 262, respectively, accounting for 2.28%, 3.84%, 15.96% and 22.13% of their respective total numbers of OTUs. The dilution curves of OTUs tended to be gradual, indicating that the measured data could accurately reflect the information on the plant fungal community (Fig. 2, c, d).

Table 1 shows that the number of Observed_species, Chao1 index and ACE number index were highest in rhizosphere soil, followed by rhizosheath soil, and were smallest in the root system, and there were significant differences between them. There was no significant difference in the Shannon index between rhizosphere soil and rhizosheath soil, which were both significantly higher than the Shannon index in root epidermis and roots, but the Shannon index of bacteria in root epidermis was significantly higher than in roots. The Simpson index of bacteria in rhizosphere soil, rhizosheath soil and root epidermis had no significant differences, but were significantly higher than those in the root system. The goods_coverage index of the samples was higher than 98%.

The observed_species, Shannon, Simpon, Chao1, ACE, PD_whole_tree index of fungi in rhizosphere soil were the highest, followed by rhizosheath soil, but there was no significant difference between the two groups. The observed_species, Shannon, Simpon, Chao1, ACE, index of rhizosphere soil and rhizosheath soil fungi were significantly larger than the root epidermis and root. There was no significant difference in observed_species, Shannon, Simpon, Chao1, ACE index between roots and root epidermis. The good_coverage indexes of the samples were all higher than 99%.

Table 1

Alpha diversity of bacteria and fungi in roots, rhizosheath, rhizosphere and non-rhizosphere soil
Microorganism	Sample name	Observed_ species	Shannon	Simpson	Chao1	ACE	Goods_ coverage
Bacteria	SG.EC	2769.0 ± 143.9d	5.41 ± 0.31c	0.82 ± 0.03b	3210.4 ± 159.4d	3312.3 ± 174.8d	0.989 ± 0.001
	SG.RS	3287.4 ± 213.2c	8.87 ± 0.46b	0.99 ± 0.01a	3714.6 ± 248.7c	3798.1 ± 266.4c	0.989 ± 0.001
	SG.RH	3919.2 ± 187.9b	9.77 ± 0.28a	1.00 ± 0.00a	4392.5 ± 145.6b	4444.4 ± 156.3b	0.988 ± 0.000
	SG.SO	4278.6 ± 256.5a	10.10 ± 0.10a	1.00 ± 0.00a	4858.4 ± 512.0a	4938.6 ± 504.8a	0.986 ± 0.003
Fungi	SG.EC	230.2 ± 27.5b	2.38 ± 0.59b	0.62 ± 0.14b	264.3 ± 34.5b	278.1 ± 37.7b	0.999 ± 0.000
	SG.RS	223.8 ± 29.2b	2.61 ± 0.75b	0.68 ± 0.15b	260.8 ± 33.4b	272.6 ± 31.3b	0.999 ± 0.000
	SG.RH	532.2 ± 65.5a	5.11 ± 0.62a	0.89 ± 0.06a	597.7 ± 65.4a	613.4 ± 68.8a	0.998 ± 0.000
	SG.SO	563.0 ± 70.9a	5.64 ± 0.44a	0.93 ± 0.04a	693.9 ± 163.6a	656.6 ± 83.6a	0.998 ± 0.001

Note: Different lowercase letters indicate significant differences between bacterial or fungal groups (P＜0.05). Observed_ species and Goods_ coverage represents sequencing depth index; Shannon and Simpson represent the diversity index of the bacterial community, and Chao1 and ACE represent the index of bacterial community richness; SG.EC, SG.RS, SG.RH, SG.SO refer to roots, root epidermis, rhizosheath, and rhizosphere soil of Stipa grandis, respectively.

The top 10 phylum level classification of bacterial communities is shown in Fig. 3 (a). The relative abundance of Cyanobacteria in the root system was 46.4%, while the relative abundance of Cyanobacteria in rhizosphere, rhizosphere and non-rhizosphere soil was only 5.0%, 2.1% and 1.1%, respectively. Actinobacteria was the dominant population in root epidermis, and its relative abundance was 43.0%, which was significantly higher than that in the root system, rhizosheath soil and rhizosphere soil. The relative abundance of Proteobacteria in root epidermis, rhizosheath soil and rhizosphere soil was 30.9%, 31.3% and 30.2%, respectively, while that in roots was only 11.7%. The relative abundances of Acidobacteria, Gemmatimonadetes, Bacteroidetes and Verrucomicrobia in rhizosheath soil and rhizosphere soil were similar and significantly higher than those in root systems and root epidermis. The relative abundances in rhizosheath soil were 22.1%, 9.2%, 8.2% and 3.2%, respectively, and those in rhizosphere soil were 21.8%, 9.6%, 7.6% and 3.7%, respectively.

The top 10 phylum level classification of the sample fungal communities is shown in Fig. 3 (b). Different from bacterial communities, the analysis shows that Basidiomycota and Ascomycota have higher relative abundance in roots, root epidermis, rhizosheath and rhizosphere soil. The relative abundance of Basidiomycota in roots and root epidermis are 72.3% and 70.3%, respectively, which is significantly higher than in rhizosheath soil and rhizosphere soil. The relative abundance of Ascomycota in roots, rhizosheath and rhizosphere soil was 20.4%, 22.4% and 27.6%, respectively, while that in root epidermis was only 8.7%. The relative abundance of Others was in the order rhizosphere soil > rhizosheath soil > root epidermis > roots.

Results of non-metric multidimensional scaling (NMDS) analysis of bacteria in roots, root epidermis, rhizosheath and rhizosphere soil of Stipa grandis are shown in Fig. 3 (c). The microbial community structure of root epidermis, rhizosheath soil and rhizosphere soil are distinct and distant in the plot, while the distance between rhizosheath soil and rhizosphere soil is relatively close. The UPGMA cluster analysis of all samples also showed that the similarity of bacterial composition and relative abundance between rhizosheath soil and rhizosphere soil was high, and that there was a big difference in bacterial composition between soil, roots and root epidermis.

The results of NMDS analysis of fungi are shown in Fig. 3 (d). The fungal community structure of roots and root epidermis, and rhizosheath and rhizosphere soil were relatively close, while that of roots and root epidermis were relatively distinct from those of rhizosheath and rhizosphere soil. Through UPGMA cluster analysis of all the samples, in contrast to bacterial community clustering, rhizosheath soil and rhizosphere soil fungal communities can be clustered into one group, while root and root epidermis can be clustered into another group.

As shown in Table 2, the bacterial community composition in rhizosheath soil and rhizosphere soil of Stipa grandis was not significantly different (P > 0.05). The bacterial community composition of roots and root epidermis of Stipa grandis was significantly different at P < 0.05, and the differences in bacterial community among other groups was extremely significantly different (P < 0.01).

There was no significant difference in fungal communities between the roots and root epidermis of Stipa grandis or between rhizosheath soil and rhizosphere soil (P > 0.05). The difference of fungal community among other groups reached an extremely significant level (P < 0.01).

Table 2

Significance test table of difference in bacterial and fungal community structure between groups
Microorganism	vs_group	SS	df	MS	Fs	p-value
Bacteria	SG.EC-SG.RH-SG.RS-SG.SO	2.18148(0.234448)	3(16)	0.727159(0.014653)	49.6253	< 0.001**
	SG.EC-SG.RS	1.12673(0.0835403)	1(8)	1.12673(0.0104425)	107.898	0.013*
	SG.RS-SG.SO	0.305587(0.11343)	1(8)	0.305587(0.0141787)	21.5525	0.003**
	SG.RH-SG.RS	0.286011(0.154591)	1(8)	0.286011(0.0193239)	14.8009	0.005**
	SG.EC-SG.RH	1.28529(0.121018)	1(8)	1.28529(0.0151273)	84.9651	0.007**
	SG.RH-SG.SO	0.0135499(0.150908)	1(8)	0.0135499(0.0188635)	0.718314	0.672
	SG.EC-SG.SO	1.34579(0.0798571)	1(8)	1.34579(0.00998214)	134.82	0.006**
Fungi	SG.EC-SG.RH-SG.RS-SG.SO	6.63178(6.53519)	3(16)	2.21059(0.408449)	5.41216	< 0.001**
	SG.EC-SG.RS	0.407402(2.75986)	1(8)	0.407402(0.344982)	1.18094	0.309
	SG.RS-SG.SO	3.31283(3.25717)	1(8)	3.31283(0.407146)	8.1367	0.006**
	SG.RH-SG.RS	2.17638(3.47909)	1(8)	2.17638(0.434886)	5.00449	0.002**
	SG.EC-SG.RH	2.86233(3.27802)	1(8)	2.86233(0.409753)	6.98551	0.006**
	SG.RH-SG.SO	0.429087(3.77533)	1(8)	0.429087(0.471917)	0.909244	0.541
	SG.EC-SG.SO	4.07552(3.0561)	1(8)	4.07552(0.382013)	10.6685	0.008**

Note: * Indicates significant difference between groups (P＜0.05); * * indicates extremely significant difference between groups (P＜0.01). SS stands for total variance, also known as the sum of squares of deviation; df is degrees of freedom; MS is the mean square (difference), i.e. SS / DF; FS is the F test value; P-value is the p value, with a value less than 0.05 indicating significant difference between groups. The values corresponding to the residual items are in brackets. SG.EC, SG.RS, SG.RH, SG.SO refer to roots, root epidermis, rhizosheath, and rhizosphere soil of Stipa grandis, respectively.

Rhizoplane refers to the outer surface of plant roots and any closely attached soil or debris particles, which was proposed by Clark (1949). However, some scholars after him have different views, that is, only the soil attached to the root can be regarded as the rhizosphere, while the root epidermis washed by soil particles is called rhizosplane (Cook and Lochhead, 1959; Wieland et al. 2001; Bulgarelli et al. 2012). York et al. (2016) suggest that it was incorrect to call root epidermis as rhizoplane, which would greatly reduce the spatial range of rhizosphere and he agreed with Clark's (1949) definition of rhizosplane. The results showed that there were significant differences in the composition of bacterial and fungal communities in root epidermis and rhizosheath, which could be inferred that they played different but important roles in plant growth. We agree with Clark (1949) and York et al (2016) that the combination of rhizosheath and root epidermis is called rhizoplane and belongs to part of rhizosphere. The unique environment of the internal roots of plants is called the inner boundary of roots (compat et al. 2010), which is also recognized by York et al (2016). Our results showed that the bacterial and fungal community structure of root was significantly different from that of rhizosheath soil and rhizosphere soil, and although there was no significant difference in fungal community between root and root epidermis, there were differences in bacterial community. In addition, the total relative abundance of Actinobacteria and Proteobacteria in root epidermis was 73.9%, which was significantly different from that in root and soil. This indicates that there are different microbial community systems in root, root epidermis, rhizosheath and rhizosphere soil of rhizosheath plants, and our method of rhizosheath separation is feasible. Both sides of root epidermis are root system and rhizosheath, which can be used as a compartment to separate root system from soil. In terms of rhizosphere microbial ecology, Philippot (2013) mentioned that rhizosphere is the interface between plant roots and soil, and the rhizosphere environment is complex and dynamic. The interaction between various microorganisms affects plant growth and tolerance to biotic and abiotic stresses. However, we suggest that for plants with rhizosheath, the outer root epidermis is the interface between the root system and the external soil.

In recent years, research on the structure and function of plant roots, rhizosphere microbial communities has gradually increased (Berendsen et al. 2012; Tkacz et al. 2015; Shi et al. 2016). Research has not only focused on changes in microbial diversity, but also included study of several different interactions and mechanisms between plant and soil microorganisms. For example, a recent review mentioned that the interaction of underground root soil microorganisms is very important for the growth and health of aboveground plants, and discussed the overall view of root soil rhizome microbial interaction realized by the progress of omics and bioinformatics technology, and the potential strategies for managing complex rhizosphere interaction to improve crop yield (Lambers et al. 2009; Zhang et al. 2015; Dessaux et al. 2016). Studies on roots, rhizosheath, soil and microorganisms and the mechanisms of their interaction are rare. The main reasons may include: (1) Not all plants have a rhizosheath, and its function may be underestimated; (2) there is no standard method for separation of the rhizosheath from rhizosplane.

In this study, phosphate buffer was used as a protective substance to separate roots, root epidermis, rhizosheath, and rhizosphere soil, which seems to be able to clarify the relationship between rhizosheath, root epidermis and microbial differences in root. Bergmann et al (2009) found that the growth of several Gramineae plants in low nutrient and low moisture dune environments may be mediated by nitrogen fixing bacteria related to the rhizosheath. In addition, studies have shown that the rhizosheath can improve the drought resistance of plants (Pate and Dixon, 1996; Shane et al. 2010; Benard et al. 2016), but whether this is related to the function of some microorganisms in the rhizosheath needs further experiments to prove. It has been proved that there are nitrogen fixing bacteria in the rhizosheath, and nitrogen fixing bacteria can provide ammonia to plants in nitrogen limited soil (Wullstein et al. 1979;Wullstein 1980༛Buckley 1982). There are few studies on other functions of microorganisms in root epidermis, which may be limited by experimental techniques. The differences in composition and function of microorganisms inside and outside the rhizosheath may prompt us to reexamine its potentially important role in plant growth. This study will provide a new method and theoretical guidance for further exploring the function and ecological significance of rhizosheath.

This study provides a feasible method to separate rhizosheath and root epidermis. This will provide the possibility for further study on the microorganism of rhizosheath and root surface. There are significant differences in bacterial diversity between root epidermis and roots, rhizosheath and rhizosphere soils, suggesting that some functions of root epidermis and rhizosheath may play an important but previously ignored role. We suggest that the root epidermis can be used as a compartment to separate the root system from the soil and that it may act as the interface between the host and the external environment, with microorganisms in this interface potentially having some important functions.

SG.EC: roots; SG.RS: root epidermis; SG.RH: rhizosheath; SG.SO: rhizosphere soil.

Ethics approval and consent to participate

Not applicable

Consent for publication

Not applicable

Availability of data and materials

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

Competing interests

All authors declare that they have no competing interests.

Funding

This work was supported by The National Natural Science Foundation of China

(Grant No. 31760146), Key Laboratories of Ministry of Education (IRT-17R59). The funding body had no role in the design of the study and collection, analysis, and interpretation of data and in writing the manuscript.

Authors’ contributions

Ai-min Zhu was responsible for writing the paper; Guo-dong Han were responsible for the design and management of the experiment. Hai-li Liu, Yue-hua Wang assisted Ai-min Zhu in sampling and DNA extraction.

Acknowledgements

We are very grateful to Mr. Cheng Xu (Wageningen University,) for his help in extracting microbial DNA from plants and soil.

Authors’ information

¹Key Laboratory of Grassland Resources, Ministry of Education. College of Grassland, Resources and Environment; Key Laboratory of Forage Cultivation, Processing and High Efficient Utilization of the Ministry of Agriculture; Inner Mongolia Key Laboratory of Grassland Management and Utilization, Inner Mongolia Agricultural University, Hohhot, Inner Mongolia, 010019, China

Bailey C, Scholes M (1997) Rhizosheath occurrence in south African grasses. S Afr J Bot 63:484–490. https://doi.org/10.1016/S0254-6299(15)30803-6
Bergmann D, Zehfus M, Zierer L, Smith B, Gabel M (2009) Grass Rhizosheaths: Associated bacterial communities and potential for nitrogen fixation. Western North American Naturalist 69(1):105–114. https://doi.org/10.3398/064.069.0102
Berendsen RL, Pieterse CM, Bakker PA (2012) The rhizosphere microbiome and plant health. Trends Plant Sci 17:478–486. https://doi.org/10.1016/j.tplants.2012.04.001
Bokulich NA, Subramanian S, Faith JJ, Gevers D, Gordon JI, Knight R, Mills DA, Caporaso JG (2013) Quality-filtering vastly improves diversity estimates from Illumina amplicon sequencing[J]. Nat Methods 10(1):57–59. https://doi.org/10.1038/nmeth.2276
Brown LK, George TS, Neugebauer K, White PJ (2017) The rhizosheath – a potential trait for future agricultural sustainability occurs in orders throughout the angiosperms. Plant Soil 418(1–2):115–128. https://doi.org/10.1007/s11104-017-3220-2
Buckley R (1982) Sand rhizosheath of an arid zone grass. Plant Soil 66:417–421. https://doi.org/10.1007/BF02183809
Benard P, Kroener E, Vontobel P, Kaestner A, Carminati A (2016) Water percolation through the root-soil interface. Adv Water Res 95:190–198. https://doi.org/10.1016/j.advwatres.2015.09.014
Caporaso JG, Kuczynski J, Stombaugh J, Bittinger K, Bushman FD, Costello EK, Fierer N, Pena AG, Goodrich JK, Gordon JI (2010) QIIME allows analysis of high-throughput community sequencing data. Nat Methods 7(5):335–336. https://doi.org/10.1038/nmeth.f.303
Clark FE (1949) Soil microorganisms and plant roots. Adv Agron 1:241–288. https://doi.org/10.1016/S0065-2113(08)60750-6
Pieterse, Corne MJ, Jonge D, Ronnie, Berendsen, Roeland L (2016) The Soil-Borne Supremacy. Trends in Plant Science
Dessaux Y, Grandclement C, Faure D (2016) Engineering the rhizosphere. Trends Plant Sci 21:266–278. https://doi.org/10.1016/j.tplants.2016.01.002
Delhaize E, Rathjen TM, Cavanagh CR (2015) The genetics of rhizosheath size in a multiparent mapping population of wheat. J Exp Bot 66:4527–4536. https://doi.org/10.1093/jxb/erv223
Duell RW, Peacock GR (1985) Rhizosheaths on mesophytic grasses. Crop Sci 25:880–883. https://doi.org/10.2135/cropsci1985.0011183X002500050036x
Edgar RC (2013) UPARSE: highly accurate OTU sequences from microbial amplicon reads. Nat Methods 10(10):996–998. https://doi.org/10.1038/NMETH.2604
Haas BJ, Gevers D, Earl AM, Feldgarden M, Ward DV, Giannoukos G, Ciulla D, Tabbaa D, Highlander SK, Sodergren E (2011) Chimeric 16S rRNA sequence formation and detection in Sanger and 454-pyrosequenced PCR amplicons. Genome research 21(3):494–504. https://doi.org/10.1101/gr.112730.110
George TS, Brown LK, Ramsay L, White PJ, Newton AC, Bengough AG,RussellJ, Thomas WTB (2014) Understanding the genetic control and physiological traits associated with rhizosheath production in barley (Hordeum vulgare). New Phytol 203:195–205. https://doi.org/10.1111/nph.12786
Goodchild DJ, Myers LF (1987) Rhizosheaths - a neglected phenomenon in Australian agriculture. Aust J Agric Res 38:559–563. https://doi.org/10.1071/ar9870559
Jenny H, Grossenbacher K (1963) Root-soil boundary zones as seen in the electron microscope. Soil Sci Soc Am J 27:273–277. https://doi.org/10.2136/sssaj1963.03615995002700030018x
Othman AA, Amer WM, Fayez M, Hegazi NA (2004) Rhizosheath of Sinai desert plants is a potential repository for associative diazotrophs. Microbiol Res 159:285–293. https://doi.org/10.1016/j.micres.2004.05.004
Pate JS, Dixon KW (1996) Convergence and divergence in the southwestern Australian flora in adaptations of roots to limited availability of water and nutrients, fire and heat stress, New South Wales, pp 249–258
Philippot L, Raaijmakers JM, Lemanceau P, Vander PWH (2013) Going back to the roots: the microbial ecology of the rhizosphere. Nature Reviews Microbiology, 2013, 11(11):789–799. https://doi.org/10.1038/nrmicro3109
Pohlman AA, Mccoll JG (1982) Nitrogen fixation in the rhizosphere and rhizoplane of barley. Plant Soil 69:341–352. https://doi.org/10.1007/BF02372455
Price SR (1911) The roots of some north African desert-grasses. New Phytol 10:328–340
Quast C, Pruesse E, Yilmaz P, Gerken J, Glckner FO (2012) The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucl Acids Res, 41(D1). https://doi.org/10.1093/nar/gks1219
Lambers H, Mougel C, Jaillard B, Hinsinger (2009) Plant–microbe–soil interactions in the rhizosphere: an evolutionary perspective. Plant Soil 321:83–115. https://doi.org/10.1007/s11104-009-0042-x
Lynch JP (2007) Roots of the second green revolution. Aust J Bot 55:493–512. https://doi.org/10.1016/S0264-410X(02)00552-2
Magoč T, Salzberg SL (2011) FLASH: fast length adjustment of short reads to improve genome assemblies. Bioinformatics 27(21):2957–2963. https://doi.org/10.1093/bioinformatics/btr507
McCully ME (2003) Roots in soil: unearthing the complexities of roots and their rhizospheres. Annu Rev Plant Phys 50:695–718. https://doi.org/10.1146/annurev.arplant.50.1.695
Shane MW, McCully ME, Canny MJ, Pate JS (2011) Development and persistence of sandsheaths of Lyginia barbata (Restionaceae): relation to root structural development and longevity. Ann Bot 108:1307–1322. https://doi.org/10.1093/aob/mcr244
Shane MW, McCully ME, Canny MJ, Pate JS, Hai N, Mathesius U, Cawthray GR,Lambers H (2009) Summer dormancy and winter growth: root survival strategy in a perennial monocotyledon. New Phytol 183:1085–1096. https://doi.org/10.1136/ard.2008.105957
Shane MW, McCully ME, Canny MJ, Pate JS, Huang C, Ngo H, Lambers H (2010) Seasonal water relations of Lyginia barbata (southern rush) in relation to root xylem development and summer dormancy of root apices. New Phytol 185:1025–1037. https://doi.org/10.1111/j.1469-8137.2009.03143.x
Shi S, Nuccio EE, Shi ZJ, He Z, Firestone MK (2016) The interconnected rhizosphere: high network complexity dominates rhizosphere assemblages. Ecol Lett 19:926–936. https://doi.org/10.1111/ele.12630
Spaepen S, Bossuyt S, Vanderleyden J, Engelen K, Marchal K (2014) Phenotypical and molecular responses of Arabidopsis thaliana roots as a result of inoculation with the auxin-producing bacterium Azospirillum brasilense. New Phytologist, 201
Sprent JI (1975) Adherence of sand particles to soybean roots under water stress. New Phytol 74:461–463
Tkacz A, Cheema J, Chandra G, Grant A, Poole PS (2015) Stability and succession of the rhizosphere microbiota depends upon plant type and soil composition. The ISME Journal 9:2349–2359. https://doi.org/10.1038/ismej.2015.41
Unno Y, Okubo K, Wasaki J, Shinano T, Osaki M (2005) Plant growth promotion abilities and microscale bacterial dynamics in the rhizosphere of lupin analysed by phytate utilization ability. Environ Microbiol 7:396–404. https://doi.org/10.1111/j.1462-2920.2004.00701.x
Volkens G (1887) Die Flora der ægyptisch-arabischen Wuste auf Grundlage anatomisch-physiologischer Forschungen. Gerbruger Borntraeger, Berlin, p 156
Vries FTD, Griffiths RI, Knight CG, Nicolitch O, Williams A (2020) Harnessing rhizosphere microbiomes for drought-resilient crop production. Science, 368. https://doi.org/10.1126/science.aaz5192
Wang Q (2007) Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl Environ Microbiol 73:5261–5267
Wullstein LH (1980) Nitrogen fixation (acetylene reduction) associated with rhizosheaths of Indian rice–grass used in stabilization of the Slick Rock, Colorado tailings pile. J Range Manag 33:204–206. https://doi.org/10.2307/3898285
Wullstein LH, Bruening ML, Bollen WB (1979) Fixation associated with sand grain root sheaths (rhizosheaths) of certain Xeric grasses. Physiol Plant 46:1–4. https://doi.org/10.1111/j.1399-3054.1979.tb03176.x
York LM, Carminati A, Mooney SJ, Ritz K, Bennett MJ (2016) The holistic rhizosphere: integrating zones, processes, and semantics in the soil influenced by roots. J Exp Bot 67(12):3629–3643. https://doi.org/10.1093/jxb/erw108
Young IM (1995) Variation in moisture contents between bulk soil and the rhizosheath of wheat (Triticum aestivum L. cv. Wembly) New Phytologist 130:125–139
Zhang R, Vivanco JM, Shen Q (2017) The unseen rhizosphere root–soil–microbe interactions for crop production. Curr Opin Microbiol 37:8. https://doi.org/10.1016/j.mib.2017.03.008
Zhang Y, Ruyter-Spira C, Bouwmeester HJ (2015) Engineering the plant rhizosphere. Curr Opin Biotechnol 32:136–142. https://doi.org/10.1016/j.copbio.2014.12.006

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Analysis of Microbial Community Structure Around Roots of Stipa Grandis

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Abstract

Figures

Introduction

Materials And Methods

Overview of the research site

Test sample collection

Harvest of roots, root epidermis, rhizosheath, rhizosphere

Sequencing process (Novogene company)

Data analysis based on the Analysis Platform of Novogene company

Results And Analysis

Distribution characteristics of bacterial and fungal communities

Discussion

Conclusion

Abbreviations

Declarations

References

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