Spatiotemporal dynamics of endophytic fungal diversity in the roots of Amomum villosum Lour

doi:10.21203/rs.3.rs-4995227/v1

Download PDF

Article

Spatiotemporal dynamics of endophytic fungal diversity in the roots of Amomum villosum Lour

https://doi.org/10.21203/rs.3.rs-4995227/v1

This work is licensed under a CC BY 4.0 License

You are reading this latest preprint version

This study aims to investigate the colonization and diversity of endophytic fungi in Amomum villosum roots under different planting locations and growth ages and to analyze the fungal composition. In this study, we performed Illumina-based ITS rDNA sequencing to investigate the effects of growth ages, and sample plots on the rhizosphere fungi of A. villosum. The results of the colonization analysis showed that arbuscular mycorrhizal fungi (AMF) and dark septate endophytes (DSEs) widely colonized the roots of A. villosum, but the colonization abundance no longer increased significantly with increasing growth age. Glomus was the dominant AMF, and Exophiala, Cladosporium and Cladophialophora were the dominant DSEs. Phoma, Acremonium, Myrothecium and Trichoderma were supposed to be the dominant beneficial fungi. Alpha and beta diversity showed that the diversity, abundance and community composition of root fungi were not significantly correlated with growth ages but were affected by planting locations. Taken together, the diversity and abundance of fungal communities in the roots differed significantly by planting location, and some were affected by the growth ages of A. villosum. DSEs were considered to be dominant beneficial microorganisms and were largely responsible for the growth and development of A. villosum, weakening continuous cropping obstacles.

Biological sciences/Microbiology

Biological sciences/Microbiology/Fungi

Biological sciences/Plant sciences

Biological sciences/Plant sciences/Plant physiology

Amomum villosum Lour.

endophytic fungi (EF)

colonization rate

fungal community composition

dark septate endophyte (DSE

arbuscular mycorrhizal fungi (AMF)

Microorganisms are an inseparable part of plant interactions during long-term growth. Healthy plants are rich in microbial resources both inside and outside of roots, stems and leaves, which comprise a large plant microbiota [1, 2]. As a major vehicle for soil microorganisms to colonize plants, plant roots gather plenty of endophytic fungi (EF), e.g., Ascomycota, Basidiomycota and Glomeromycota [3]. Arbuscular mycorrhizal fungi (AMF) and dark septate endophytes (DSEs) are the two main groups of EF that play important roles in nutrient uptake and resistance to biotic or abiotic stresses. Both of them are capable of forming mycorrhizal symbioses in root cells [4, 5]. Symbiosis is based on the acquisition of carbon sources from the host plant, which provides the feedback on important nutrients to plants by regulating the release of signaling molecules, e.g., transporter proteins, bioactive substances and Myc factors [6, 7]. Numerous studies have shown that mycorrhizal symbioses directly improve the resistance of host plants to stresses of drought, salt, heavy metal and low phosphorus, but these mechanisms of action need to be further refined [8–11]. The interactions of microbe‒microbe and microbe‒plant interactions are complex and inevitable, and they are what we must study and analyze in depth, which will be helpful to understand the structure of the microbial‒plant interactions in nature.

Root endophytic fungal community composition is diverse and complex and is often affected by the health status, developmental stage and growth environment of the plant. Huang et al. found that although the root fungi of healthy and diseased lisianthus had similar fungal communities, healthy plants had more potentially disease-resistant fungi, while diseased plants were enriched in more pathogenic fungi [12]. In studies by Šmilauer et al. and Kil et al., the diversity of AMF in the roots of lower-age plants was found to be significantly lower than that in higher-age plants [13, 14]. In addition, both nutrient contents in the soil and its geographical location are also key factors that significantly affect the root endophytic fungal community structure [11, 15].

Isolation of EF using pure culture is an effective method for fungal diversity studies, but often the isolation and investigation of some strains are difficult to achieve due to unculturable characteristics, e.g., AMF [16, 17]. The application of Illumina sequencing technology of the internal transcribed spacer (ITS) region to accurately analyze the community composition and diversity of EFs at the molecular level can both reveal unculturable fungal species and allow us to directly understand fungal community relationships and address questions related to ecosystem function [18, 19].

Amomum villosum Lour. is a perennial medicinal herb of the genus Amomum in the Zingiberaceae family, with records of its medicinal use dating back to approximately 1,400 years [20]. The fruits have the ability to enliven the spleen, resolve dampness and harmonize the stomach, prevent abortion and regulate the intestinal microflora [21]. In China, A. villosum is mainly distributed in shady and humid mountainous areas or natural forests in Yunnan Province, Guangdong Province and Hainan Province[20]. Previous studies have indicated that the composition of A. villosum rhizosphere soil bacteria and fungi is complex and diverse and presumed to be correlated with its growth and fruit yield and quality [23]. However, there are no systematic studies on root EFs that form a symbiotic relationship with A. villosum, and the community composition and abundance are unknown. Therefore, it is urgent to study the EF of A. villosum roots.

Colonization investigation and molecular measurements of A. villosum root EFs and thus a comprehensive assessment are necessary to broaden the understanding of microbial diversity. In this study, our objectives were to investigate the fungal community structure and diversity of A. villosum using high-throughput sequencing techniques and thereafter uncover the changes associated with different planting locations and growth ages.

2.1 Sample collection

As shown in Fig. 1a, plant samples were collected from Yunnan Province and Guangdong Province, which are the main production regions of A. villosum across southern China. Plant samples were mainly sampled from Xishuangbanna plots (BNP), Pu'er plots (PEP), Honghe plots (HHP) and Wenshan plots (WSP) of Yunnan Province, and Yangchun plots (YCP) of Guangdong Province in September 2021 (Fig. 1a). A. villosum samples consisted of two age groups of one and three years old, which were newly sprouted plants in 2020 and planted in 2018. In total, ten group samples, including BNP-1, BNP-3, PEP-1, PEP-3, HH-1, HH-3, WSP-1, WSP-3, YCP-1 and YCP-3, were collected for analysis. The two groups of one- and three-year-old plants in each plot were collected by a five-point sampling method (Fig. 1b); that is, there were ten plants of each age group in each plot. Fresh fibrous roots of A. villosum were collected and transported to the laboratory and washed repeatedly with sterile water to remove visible soil particles. The fibrous roots of ten samples for each group were mixed well for a total of three biological replicates and placed in 50 mL sterile EP tubes in two portions. One portion was sent to Ronin Bio (China) for high-throughput sequencing, and the other was stored at -80°C for further analysis [24].

2.2 Assessment of fungal colonization in roots

All fibrous roots that were potentially colonized were cleaned thoroughly and stained. Afterward, they were squashed and mounted in lactic acid-glycerol on microscope slides. Root endophyte quantification was performed by the magnified intersection method using a compound microscope (Nikon, Eclipse E100), magnified × 40. Roots of fixed length (1.5 cm) were scored for approximately 100 fields of view. The colonization rates of AMF and DSEs were generated by counting the number of view fields of typical structures, e.g., vesicles, arbuscules and hyphal circles for AMF and dark mycelium, septate hyphae and microsclerotias for DSEs[8].

2.3 DNA extraction, PCR amplification and sequencing

Genomic DNA of the root fungi was extracted and purified using the Zymo Research BIOMICS DNA Microprep Kit (Zymo Research, USA) according to the manufacturer's protocol. The ITS rDNA sequencing of fungi was amplified with primers ITS3 (5'-GATGAAGAACGYAGYRAA-3') and ITS4 (5'- TCCTCCG CTTATTGATATGC-3') by polymerase chain reaction (PCR) [25]. Each sample was amplified in triplicate in a 25 µL system containing 1 U KOD-Plus-Neo polymerase, 1x PCR Buffer for KOD-Plus-Neo, 0.2 mM dNTPs 1.5 mM MgSO4 and 0.3 µM forward and reverse primers, 20 ng/µL template DNA, and GSS Depletion Mix (Rhonin Bio, China) to remove interference from the elimination of chloroplast and mitochondrial sequences in the samples. Amplification was performed with an Applied Biosystems® PCR System 9700 (USA) according to the following criteria: 94°C/1 min for one cycle; 94°C/20 s, 54°C/30 s and 72°C/30 s for 25–30 cycles; and 72°C/5 min for one cycle. Afterward, the PCR products were analyzed by electrophoresis on 2% agarose gels and purified and detected for quantification.

2.4 cDNA library construction and sequencing

Library construction was performed using the NEBNext Ultra II DNA Library Prep Kit for Illumina (BioLabs Inc., New England). After that, purified amplicons were pooled in equimolar ratios and paired-end sequenced on an Illumina MiSeq PE250 platform (Illumina, San Diego, USA) according to the standard protocols by Ronin Bio (China).

2.5 Data processing and community composition analysis

Spliced double-ended sequences were obtained as raw tags. Then, quality filtering was conducted using the tool of Quantitative Insights Into Microbial Ecology (QIIME, v1.8.0, http://qiime.org/) to obtain high-quality clean tags, and the chimeras were removed using the Uchime algorithm [26, 27]. Based on the Usearch software (http://drive.com/uparse/), OTU clustering was performed using the UPARSE algorithm at a 97% sequence identity [28]. The sequence with the highest frequency of occurrence in each OTU was selected as a representative sequence for the OTU and annotated for the analysis using the UCLUST taxonomy with the SILVA database [29]. Subsequently, data transformation was performed using R software (Version 2.15.3), and graphical visualization of community composition was then completed with the ggplot2 package in R.

The alpha diversity indices (Chao1, ACE, Shannon, Simpson, Good's coverage) were calculated using the R software Vegan package, and the PD indices were generated by the Picante package [30]. The differences in parameter variance among samples from different sample plots were estimated by ANOVA. Multiple comparisons were performed using the agricolae package. Differences between annual and triennial samples were tested using the Wilcoxon rank sum test (Wilcox. test, stats package). Main coordinates analysis (PCoA, GuniFrac package in R language) was performed using a weighted UniFrac distance index to detect similarity between samples [31]. To analyze differences in community composition, PERMANOVA (adonis function, vegan package) was performed to explain the significance of temporal and spatial effects on sample differences at a significance level of p < 0.05. Key genera between groups were identified by comparison with Random forest analysis and abundance difference test analysis.

3.1 Colonization rates of AMF and DSE in A. villosum roots

Typical structures of AMF and DSEs were widely observed in the intracellular or intracellular space of A. villosum roots, including hyphae, vesicles and arbuscules for AMF and microsclerotia and septate hyphae for DSEs (Fig. 2). Hyphae, arbuscules, microsclerotia and septate hyphae are signal that the symbiotic plant‒AMF/DSE system is forming, thereafter exchanging carbon sources and energy. High colonization rates of AMF in annual samples presented at sample plots of HHP, WSP and PEP with values of 34.29 ± 5.02, 32.39 ± 8.72 and 28.72 ± 5.07%, respectively, followed by 10.07 ± 4.74% of YCP and 3.87 ± 3.38% of BNP (Table S1). The AMF colonization rates of the BNP, WSP and YCP samples sharply increased as the growth age increased three years, especially for the triennial sample YCP, with an increased rate of 251.44%. For the DSE, colonization rates for all samples were lower than those of AMF. Annual PEP samples had the largest DSE colonization rate of 9.14 ± 4.73%, but no typical structures were observed in the annual BNP sample. Compared with annual samples, the colonization rates of triennial samples BNP, HHP and YCP increased significantly, while samples of WSP and PEP decreased or presented the no significantly changes.

3.2 Sequencing results

A total of 985,690 high-quality sequences were obtained from all samples (31,127 ~ 38,943 reads per sample). After UCHIME analysis of the samples to remove chimeric and somatic sequences, effective tags were obtained for subsequent analysis with an average length of 341 bp, in the range of 336 ~ 352 bp. Sequences were then aggregated into 1,092 fungal operational taxonomic units (OTUs) with a 97% identity threshold. All sequences were annotated to 9 phyla, 26 classes, 69 orders, 149 families, 254 genera and 270 species.

3.3 Composition of endophytic fungal communities

Venn diagrams showed that 333 OTUs were shared between annual and triennial samples, accounting for a proportion of 30.49% (Fig. 3a). There were 429 OTUs specific to one year and 330 OTUs specific to three years, which accounted for 39.29% and 30.22% of the total OTU richness, respectively, suggesting that the diversity of EF in A. villosum roots decreased with increasing growth age. For the annual samples, excluding 72 common OTUs, the OTUs specific to each group were ranked as HHP (153) > YCP (132) > WSP (125) > WSP (102) > BNP (58) (Fig. 3b), and the sort order was BNP (130) > YCP (110) > HHP (103) > WSP (98) > PEP (70) for triennial samples without 71 common OTUs (Fig. 3c). As shown in Fig. 3d, the rarefaction curves of fungal OTUs showed that the HHP annual samples had higher OTUs than those of all samples, and the annual samples of BNP had the lowest number of OTUs.

As shown in Fig. 4, Ascomycota was the predominant phylum for annual and triennial samples with ratios of 93.58 and 92.39%, respectively, followed by Basidiomycota (2.96 and 5.75%). Glomeromycota was another main phylum with ratios of 0.77/0.22% (one-year/three-year), which belongs to the taxon of AMF. The common dominant class was Sordariomycetes (81.88 and 71.77%%, one-year/three-year), in which Dothideomycetes accounted for 7.61% more in the triennial samples than in the annual samples. Pleosporales (13.00 and 19.53%, one-year/three-year) and ectomycorrhizal fungi (EMF) Hypocreales (66.63 and 50.53%, one-year/three-year) were the common dominant orders in annual and triennial samples, respectively (Figure S1). Cordycipitaceae (51.66 and 15.92%, 1-year/3-year) and Nectriaceae (8.28 and 7.15%, one-year/three-year) were the codominant families (Figure S1). In addition, compared to 0.21% of annual samples, the ratio of the family Cladosporiaceae had a significant increase in triennial samples and was up to 3.15%, which was usually supposed to be a growth-promoting microorganism.

At the genus level, the top 20 abundant genera were selected to generate mapping images separately (Fig. 5). The EMF genera Beauveria (51.47/35.05%, one-year/three-year) and Fusarium (6.43/8.52%, one-year/three-year) were identified as the main codominant genera in annual and triennial samples. The potential antimicrobial fungi Phoma (1.65/4.32%, one-year/three-year), the saprophytic fungi Acremonium (3.87/2.24%, one-year/three-year) and Myrothecium (0.09/0.02%, one-year/three-year), and the biocontrol fungi Trichoderma (0.88/1.04%, one-year/three-year) were major genera for all samples in this study. The proportions of Exophiala, Cladosporium and Cladophialophora showed an increasing trend with ranges of 0.22 ~ 1.18, 0.18 ~ 3.05 and 0.15 ~ 0.25%, respectively, of which these genera could be classified as DSEs in taxonomic status. It is speculated that these fungi play an important role in the succession of A. villosum. In addition, Glomus (0.12/0.04%, one-year/three-year) was the dominant genus of AMF but ranked below 20 in abundance. The EMF fungus Tomentella and DSE Phialocephala were found only in annual samples, and the saprophytic fungus Oidiodendron was found alone in triennial samples.

3.4 The response of the alpha diversity of root EFs to temporal space

Community richness and diversity were analyzed using five alpha diversity indices of Chao1, Shannon’s, Simpson’s, ACE and Goods coverage, and the depth index (Goods coverage) of each sample library was over 99% (99.8‒100%), indicating that the sampling was reasonable (Table. S2). The Chao1 and ACE indices are indicative of fungal community richness, and Shannon’s and Simpson’s index represent fungal community diversity. The fungal community richness and diversity of the annual root samples were not significantly different from those of the triennial samples (Fig. 6a). Among the samples from the 5 sample plots, the annual and triennial samples of the HHP plots had the highest richness and diversity and showed a significant decrease in diversity and richness after three years of growth. In contrast, the diversity and richness of the PEP, BNP and WSP samples increased significantly with increasing growth age (Fig. 6b, c).

3.5 Response of beta diversity of root EF to temporal space

Weighted UniFrac PCoA showed that the A. villosum root EF community composition was not significantly different between samples of two growth ages but rather presented among samples of different sampling plots (Fig. 7a). Independent analyses showed significant differences in fungal community composition among the five geographically located annual samples (R = 0.6424, P = 0.0020), and triennial samples also differed significantly by geographic location (R = 0.6961, P = 0.0010) (Fig. 7b, c).

3.6 Differences in microbial communities

To determine the taxonomic fungal taxa with significant differences in abundance between annual and triennial samples, random forest analysis was used to analyze biomarkers, in which a higher Gini index indicated that the fungus was more taxonomically important between the two groups (Fig. 8). Nigrospora and Cladosporium were the main biomarkers in the annual and triennial samples, respectively. The abundance of Nigrospora significantly increased with growth ages while the abundance of Cladosporium differed significantly among groups of five triennial samples. Species of Trichoderma were significantly enriched in the annual samples, and Aspergillus species were dominant in the triennial samples. The abundance of the two genera was significantly higher than that of other strains identified from the same plot samples.

In this study, we evaluated the EF colonization of A. villosum roots under different planting locations and growth ages for the first time. High-throughput sequencing technology was employed to study the abundance, composition and distribution of root EF species.

AMF and DSEs are the two main groups of EFs and play critical roles in the growth and development of host plants. Numerous studies have shown that AMF can promote plant growth and development under different habitats and nutritional conditions, particularly by enriching the phosphorus pool in the root system [32]. It has been confirmed that DSEs also have a similar function, but more evidence is needed to further elucidate the mechanism. Studies have shown that large differences exist in the colonization rates of AMF and DSEs among plants, which are affected by climate, moisture, latitude and stress factors [33–36]. For the triennial samples, AMF colonization rates of A. villosum ranged from 13.03 to 39.48% with a mean value of 27.02%, which was closer to the Longjing tea root AMF colonization rate of 27.54% [37]. Jin et al. reported a significant 70% increase in the AMF colonization rate of Solidago canadensis under dry habitats over time [38], which was in line with our results of increased AMF colonization rates of samples of BNP, WSP and YCP. However, the AMF colonization rates of the samples HHP decreased with the age, which may be related to the competition of ecological sites with other microorganisms and the use of excessive fertilizers and pesticides.

Compared to the DSE colonization rates of 21.25% of Artemisia deversa [39], annual samples of A. villosum from five sampling plots had lower DSE colonization rates, with a range of 0 ~ 9.38% and a mean value of 3.11%. However, the mean DSE colonization rate increased with increasing growth age, reaching 6.15%. In addition, the colonization rates of AMF and DSE in sample PEP-1 were relatively higher than those in other samples, supporting the hypothesis that AMF and DSE are mutually reinforcing [32]. This result was confirmed by the studies of Della Mónica et al. and Biotic et al., which reported positive correlations between AMF and DSE colonization in Trifolium repens and Achnatherum lettermanii, and both were closely related to the utilization and absorption of phosphorus [32, 40].

In this study, the difference in fungal taxa was nonsignificant among A. villosum samples from five sampling plots, and the dominant taxa at the phylum level were Ascomycota, Basidiomycota and Glomeromycota, which was the same as the composition of root fungi of Casuarina equisetifolia [35]. At the order level, Hypocreales, Chaetothyriales and Helotiales were observed in all samples, as well as the families Cordycipitaceae and Cladosporiaceae, which were identified in other plants with biocontrol potential, e.g., Fragaria ananassa [41] and Glycine max [42]. Hypocreales are widely marketed worldwide as fungicides and Cladosporiaceae fungi have potential value in the control of white rust [43, 44]. The antimicrobial fungi Phoma, saprophytic fungi Acremonium and Myrothecium and the biocontrol fungus Trichoderma were detected as the main genera in our study, which can contribute to the improvement of rhizosphere soil microstructure. The genus Beauveria was one of the dominant microbes in A. villosum roots, which not only does not cause plant diseases but is also able to prevent insect pests and exchange nitrogen sources in the host plant [45]. Therefor, the roles of Beauveria need to be viewed dialectically. It is worth mentioning that AMF and DSE were the main components of the A. villosum root system, directly supporting the colonization rates, in which Glomus of AMF and Exophiala, Cladosporium and Cladophialophora of DSE were identified as the main genera. According to previous studies, AMF and DSE were proven to be the main fungal community in Glycyrrhiza uralensis, Ferula sinkiangensis and Tinospora cordifolia, e.g., Glomus, Diversispora and Cladosporium [46–48]. The colonization of these diverse mycorrhizal fungi may form a complex network structure in or around the root system, which plays an important role in maintaining the growth of plants and resisting diseases.

The alpha diversity results showed that there was no significant difference in richness and diversity between the annual and triennial samples, while some differences existed due to spatial differences in the samples. Similar to the results reported by Huang et al., the diversity indices of C. equisetifolia endophytic fungal communities did not differ significantly between forest ages [35]. This phenomenon may be attributed to the fact that plant root endophytic fungal diversity is largely influenced by local biotic and abiotic influences [49], e.g., soil properties as well as human disturbances both cause changes in root microbial diversity [50, 51]. It is notable that fungal diversity was consistently highest in the HHP sample plot among the five plots, which may be the result of multiple factors. However, the abundance of AMF decreased in triennial samples of HHP, suggesting that it may be related to the change in soil nutrient composition over time and may require the involvement of other microorganisms for adjustment.

The community composition analysis results showed that the fungal community composition of A. villosum did not differ significantly by age, and the spatial location was the main cause of community differences. Random forest analysis revealed that the fungal taxa were responsible for the differences in community composition to some extent. In line with the report of Zhong et al[52], Nigrospora was taken as a biomarker or key taxon in this study. It was found that the AMF strain Diversispora was involved in the growth promotion of Chrysanthemum morifolium under salt stress [53], and the strain was significantly enriched in annual samples, implying that it may play a key role in this phase. In contrast, Aspergillus and Cladosporium (DSE) were enriched in triennial samples and showed significant differences from annual A. villosum samples, of which these strains were reported to be valuable for heavy metal resistance and may be related to their involvement in maintaining and promoting plant growth [54, 55]. Colonization surveys and molecular diversity analyses responded to the importance that AMF and DSEs were involved in the evolution of plant rhizosphere fungal communities and that there are specific community structure changes due to spatial and temporal differences.

Through a fungal colonization rate survey and high-throughput sequencing, we found that a large number of fungi, mainly AMF and DSEs, adapted to environmental changes in the interrhizosphere of A. villosum. Fungal colonization rate and community structure showed differences in root systems due to the variation in location and growth ages. AMF and DSE had relatively high colonization densities in the roots of A. villosum, in which Glomus of AMF and Exophiala, Cladosporium and Cladophialophora of DSE were identified as the dominant genera, and the abundance increased with age. According to random forest analysis, Nigrospora and Cladosporium were recognized as EF biomarkers in annual and triennial samples, respectively, which may be closely linked to the growth of A. villosum. Taken together, the study provides information on the composition and diversity of fungal communities within the root system of A. villosum under different planting locations and growth ages and explores the variation in the composition of beneficial microorganisms in the root systems, which warrants further isolation and study of the role of these microorganisms in crop succession, plant growth and development and stress resistance.

Author Contributions: Guan-Hua Cao: Writing – original draft; writing – review and editing; conceptualization; data curation; formal analysis; funding acquisition; project administration; and validation; Xiao-Gang Li: Writing – original draft; writing – review and editing; data curation; formal analysis; investigation; and methodology; Xiao-Xu You: Writing – original draft; investigation; methodology; and validation; Xing-Kai Zhang: Investigation; methodology; and validation; Wen Gu: Writing – original draft; investigation; and methodology; Pei Yang: Writing – original draft; investigation and methodology; Sen He: Writing – original draft; writing – review and editing; conceptualization; data curation; formal analysis; funding acquisition; investigation; project administration; resources; and supervision; validation; Jie Yu: Writing – review and editing; conceptualization; funding acquisition; and project administration; supervision.

Funding: This research was supported by Yunnan Provincial Science and Technology Department- Applied Basic Research Joint Special Funds of Chinese Medicine (202001AZ070001-009, 202101AZ070001-014), the National Natural Science Foundation of China (82360750, 82260743), the Yunnan Fundamental Research Projects (202201AT070219, 202001AT070109), Yunnan Youth Talent Program- “Ten Thousand Plan” (YNWR-QNBJ-2020-279) and International (Foreign) Science and Technology Cooperation Base of Kunming City (GHJD-2021030).

Institutional Review Board Statement: Not applicable.

Informed Consent Statement: Not applicable.

Data Availability Statement: The data presented in this study are deposited in the National Center for Biotechnology Information (NCBI) Sequence Read Archive database (SRA) with a accession number of PRJNA1000412.

Acknowledgments: The authors are grateful to all of the laboratory members for their continuous technical advice and helpful discussions.

Conflicts of Interest: The authors declare no conflict of interest.

Horton, M.W. et al. Genome-wide association study of Arabidopsis thaliana leaf microbial community. Nat. Commun.5, 5320 (2014).
Mesny, F. et al. Genetic determinants of endophytism in the Arabidopsis root mycobiome. Nat. Commun. 12, 7227 (2021).
Rodriguez, R.J., White, J.F., Jr., Arnold, A.E. & Redman, R.S. Fungal endophytes: diversity and functional roles. New. Phytol. 182, 314-330 (2009).
van der Heijden, M.G.A., Martin, F.M., Selosse, M.A. & Sanders, I.R. Mycorrhizal ecology and evolution: the past, the present, and the future. New. Phytol.205, 1406-1423 (2015).
Jumpponen, A., Trappe, J.M. Dark septate endophytes: a review of facultative biotrophic root-colonizing fungi. New. Phytol.140, 295-310 (1998).
Bonfante, P., Genre, A. Mechanisms underlying beneficial plant-fungus interactions in mycorrhizal symbiosis. Nat. Commun. 1, 48 (2010).
Ruotsalainen, A.L. et al. Dark septate endophytes: mutualism from by-products? Trends. Plant. Sci. 27, 247-254 (2022).
Liu, C., Dai, Z., Cui, M., Lu, W. & Sun, H. Arbuscular mycorrhizal fungi alleviate boron toxicity in Puccinellia tenuiflora under the combined stresses of salt and drought. Environ. Pollut.240, 557-565 (2018).
Su, Z.Z. et al. Dark septate endophyte Falciphora oryzae-assisted alleviation of cadmium in rice. J. Hazard. Mater. 419, 126435 (2021).
Gao, M.Y. et al. Cell wall modification induced by an arbuscular mycorrhizal fungus enhanced cadmium fixation in rice root. J. Hazard. Mater. 416, 125894 (2021).
Lu, X., He, M., Ding, J. & Siemann, E. Latitudinal variation in soil biota: testing the biotic interaction hypothesis with an invasive plant and a native congener. Isme. J.12, 2811-2822 (2018).
Huang, X., Liu, S., Liu, X. & Zhang, S. Plant pathological condition is associated with fungal community succession triggered by root exudates in the plant-soil system. Siol. Biol. Biochem.151, 108046 (2020).
Šmilauer, P., Košnar, J., Kotilínek, M., Pecháčková, S. & Šmilauerová, M. Host age and surrounding vegetation affect the community and colonization rates of arbuscular mycorrhizal fungi in a temperate grassland. New. Phytol.232, 290-302 (2021).
Kil, Y.J., Eo, J.K., Lee, E.H. & Eom, A.H. Root age-dependent changes in arbuscular mycorrhizal fungal communities colonizing roots of Panax ginseng. Mycobiology42, 416-21 (2014).
Bonito, G. et al. Plant host and soil origin influence fungal and bacterial assemblages in the roots of woody plants. Mol. Ecol.23, 3356-70 (2014).
Overy, D.P., Rämä, T., Oosterhuis, R., Walker, A.K. & Pang, K.L. The neglected marine fungi, sensu stricto, and their isolation for natural products' discovery. Mar. Drugs.17 (2009).
Blau, K., Jechalke, S. & Smalla, K. Detection, isolation, and characterization of plasmids in the environment. Methods. Mol. Biol.2075, 39-60 (2020).
Nilsson, R.H. et al. Mycobiome diversity: high-throughput sequencing and identification of fungi. Nat. Rev. Microbiol.17, 95-109 (2019).
Najafi, A., Moradinasab, M., Seyedabadi, M., Haghighi, M.A. & Nabipour, I. First molecular identification of symbiotic archaea in a sponge collected from the Persian Gulf, Iran. Open. Microbiol. J.12, 323-332 (2018).
Xu, J. et al. Research on germplasm diversity of Amomum villosum. Lour in genuine producing area. PLoS One17, e0268246 (2017).
Yue, J. et al. Efficacy and mechanism of active fractions in fruit of Amomum villosum Lour. for gastric cancer. J. Cancer.12, 5991-5998 (2021).
Wang, B. et al. Effects of the continuous cropping of Amomum villosum on rhizosphere soil physicochemical properties, enzyme activities, and microbial communities. Agronomy12, 2548 (2022).
Wu, Z. et al. Temporal and spatial pattern of endophytic fungi diversity of Camellia sinensis (cv. Shu Cha Zao). BMC. Microbiol.20, 270 (2020).
Toju, H., Tanabe, A.S., Yamamoto, S. & Sato, H. High-coverage ITS primers for the DNA-based identification of ascomycetes and basidiomycetes in environmental samples. PLoS One7, e40863 (2012).
Mohsen, A., Park, J., Chen, Y.A., Kawashima, H. & Mizuguchi, K. Impact of quality trimming on the efficiency of reads joining and diversity analysis of Illumina paired-end reads in the context of QIIME1 and QIIME2 microbiome analysis frameworks. BMC. Bioinform.20, 581 (2019).
Edgar, R.C., Haas, B.J., Clemente, J.C., Quince, C. & Knight, R. UCHIME improves sensitivity and speed of chimera detection. Bioinformatics27, 2194-200 (2011).
Prodan, A. et al. Comparing bioinformatic pipelines for microbial 16S rRNA amplicon sequencing. PLoS One15, e0227434 (2020).
Edgar, R.C. Search and clustering orders of magnitude faster than BLAST. Bioinformatics26, 2460-2461 (2010).
Faith, D.P.J.B.C. Conservation evaluation and phylogenetic diversity. Biol. Conserv. 61, 1-10 (1992).
Lozupone, C., Lladser, M.E., Knights, D., Stombaugh, J. & Knight, R. UniFrac: an effective distance metric for microbial community comparison. Isme. J.5, 169-72 (2011).
Monica, I., Saparrat, M., Godeas, A.M. & Scervino, J.M.J.F.E. The co-existence between DSE and AMF symbionts affects plant P pools through P mineralization and solubilization processes. Fungal. Ecol.17, 10-17 (2015).
Lugo, M.A. et al. Arbuscular mycorrhizas and dark septate endophytes associated with grasses from the Argentine Puna. Mycologia110, 654-665 (2018).
Li, W.J. et al. Plant communities, soil carbon, and soil nitrogen properties in a successional gradient of sub-alpine meadows on the eastern Tibetan plateau of China. Environ. Manage.44, 755-65 (2009).
Huang, R. et al. Structural variability and niche differentiation of the rhizosphere and endosphere fungal microbiome of Casuarina equisetifolia at different ages. Braz. J. Microbiol.51, 1873-1884 (2020).
Veresoglou, S.D. et al. Latitudinal constraints in responsiveness of plants to arbuscular mycorrhiza: the 'sun-worshipper' hypothesis. New. Phytol.224, 552-556 (2019).
Zhang, Z. et al. Diversity in rhizospheric microbial communities in tea varieties at different locations and tapping potential beneficial microorganisms. Front. Microbiol.13, 1027444 (2022).
Jin, L., Gu, Y., Xiao, M., Chen, J. & Li, B. The history of Solidago canadensis invasion and the development of its mycorrhizal associations in newly-reclaimed land. Funct. Plant. Biol. 31, 979-986(2004).
Huo, L., Gao, R., Hou, X., Yu, X. & Yang, X. Arbuscular mycorrhizal and dark septate endophyte colonization in Artemisia roots responds differently to environmental gradients in eastern and central China. Sci. Total. Environ.795, 148808 (2021）.
Ranelli, L.B. et al. Distributions Biotic and abiotic predictors of fungal colonization in grasses of the Colorado Rockies. Divers. Distrib.21, 962-976 (2015).
Mantzoukas, S. et al. Dual action of Beauveria bassiana (Hypocreales, Cordycipitaceae) endophytic stains as biocontrol agents against sucking pests and plant growth biostimulants on melon and strawberry field plants. Microorganisms10, 2306 (2022).
Muhammad et al. Cladosporium sphaerospermum as a new plant growth-promoting endophyte from the roots of Glycine max (L.) Merr. World. J. Microb. Biot.25, 627-632 (2009).
Kepler, R.M., Maul, J.E. & Rehner, S.A. Managing the plant microbiome for biocontrol fungi: examples from Hypocreales. Curr. Opin. Microbiol.37, 48-53 (2017).
Torres, D.E. et al. Cladosporium cladosporioides and Cladosporiumpseudocladosporioides as potential new fungal antagonists of Puccinia horiana Henn., the causal agent of chrysanthemum white rust. PLoS One12, e0170782 (2017).
Barelli, L., Moonjely, S., Behie, S.W. & Bidochka, M.J. Fungi with multifunctional lifestyles: endophytic insect pathogenic fungi. Plant. Mol. Biol.90, 657-64 (2016).
Dang, H. et al. Succession of endophytic fungi and arbuscular mycorrhizal fungi associated with the growth of plant and their correlation with secondary metabolites in the roots of plants. BMC. Plant. Biol.21, 165 (2021).
Luo, Y.et al. High-throughput sequencing analysis of the rhizosphere arbuscular mycorrhizal fungi (AMF) community composition associated with Ferula sinkiangensis. BMC Microbiol.20, 335 (2020).
Mishra, A. et al. Season and tissue type affect fungal endophyte communities of the Indian medicinal plant Tinospora cordifolia more strongly than geographic location. Microb. Ecol. 64, 388-98 (2012).
Alzarhani, A.K. et al. Are drivers of root-associated fungal community structure context specific? Isme. J.13, 1330-1344 (2019).
Koczorski, P. et al. The effects of host plant genotype and environmental conditions on fungal community composition and phosphorus solubilization in willow short rotation coppice. Front. Plant. Sci.12, 647709 (2021).
Pärtel, M. et al. Historical biome distribution and recent human disturbance shape the diversity of arbuscular mycorrhizal fungi. New. Phytol. 216, 227-238 (2017).
Zhong, F. et al. Soil fungal community composition and diversity of culturable endophytic fungi from plant roots in the reclaimed area of the eastern coast of China. J. Fungi , 8, (2022).
Wang, Y., Wang, M., Li, Y., Wu, A. & Huang, J. Effects of arbuscular mycorrhizal fungi on growth and nitrogen uptake of Chrysanthemum morifolium under salt stress. PLoS One13, e0196408 (2018).
Han, S., Li, X., Amombo, E., Fu, J. & Xie, Y. Cadmium tolerance of perennial ryegrass induced by Aspergillus aculeatus. Front. Microbiol.9, 1579 (2018).
Jiang, L. et al. Plant growth promotion by two volatile organic compounds emitted from the fungus Cladosporium halotolerans NGPF1. Front. Plant. Sci. 12, 794349 (2021).

No competing interests reported.

SupplementaryMaterial202495.zip

Download PDF

Submission checks completed at journal
05 Sep, 2024
First submitted to journal
29 Aug, 2024

You are reading this latest preprint version

Spatiotemporal dynamics of endophytic fungal diversity in the roots of Amomum villosum Lour

Status:

Version 1

Abstract

Figures

1. Introduction

2. Materials and Methods

2.1 Sample collection

2.2 Assessment of fungal colonization in roots

2.3 DNA extraction, PCR amplification and sequencing

2.4 cDNA library construction and sequencing

2.5 Data processing and community composition analysis

3. Results

3.1 Colonization rates of AMF and DSE in A. villosum roots

3.2 Sequencing results

3.3 Composition of endophytic fungal communities

3.4 The response of the alpha diversity of root EFs to temporal space

3.5 Response of beta diversity of root EF to temporal space

3.6 Differences in microbial communities

4. Discussion

5. Conclusions

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1