Identification of Meloidogyne panyuensis (Nematoda: Meloidogynidae) infecting Orah (Citrus reticulata Blanco) and its impact on rhizosphere microbial dynamics: Guangxi, China

Sample collection

About 6.7 hectares of Orah (Citrus reticulata Blanco) orchard with trees aged 3 years and managed under unified agronomic conditions, was investigated in June 2022 in Wuming District, Nanning City, GZAR (22°58′45.92″N, 108°11′51.24″E) (Huang et al., 2022). Samples of Orah roots and rhizosphere soil were collected from two adjacent orchards with trees at different stages of growth: one healthy and robust and the other short and yellow. Three rows were randomly selected in the healthy or root-knot-infected Orah, and five plants in each row were mixed into one sample. After removing the surface soil, the rhizosphere soil samples were collected. The soil samples were stored in a cooler and transported to the laboratory. The rhizosphere soil was divided into two parts: (1) preserved at −80 °C to determine the rhizosphere soil microbial communities; and (2) air-dried to determine soil chemical properties. The healthy Orah samples were labeled as CRH, while the root-knot-infected Orah samples were labeled as CRN. Nematode egg tible water spinach (Ipomoea aquatica Forsk) in sterilized soil for propagation. Second-stage juveniles (J2s) were collected from the hatching eggs. Simultaneously, the fruits of healthy and nematode-infected Orah were collected to evaluate their mass and diameter.

Nematode extraction and identification

Female adults were selected from Orah root-knot tissue under an anatomical microscope, and a slide consisting of a 45% lactic acid solution was used to make an impression of the perineal pattern. The tail end of the nematode was cut with a scalpel, and the eggs and other adhesions attached to the inside were carefully removed and slightly modified with a brush, leaving only the perineal pattern part. The perineal pattern was transferred to a glass slide with one drop of pure glycerol, covered with a cover glass, and photographed under an optical microscope (Yang et al., 2021).

One single clean J2 nematode was placed into a 500 μL centrifuge tube, and 8 µL ddH2O and 1 µL 10× PCR buffer were added. The tube was placed in liquid nitrogen for 1 min, then heated to 85 °C for 2 min, and this process was repeated 7–8 times. Then, 1 µL 1 mg/mL proteinase K was added, followed by incubation at 56 °C for 1 h and 95 °C for 10 min. This nematode genomic DNA was directly used for PCR or stored at −20 °C. The rDNA-ITS sequence of the root-knot nematodes was amplified using the universal primers: V5367/26S (5′-TTGATTACGTCCCTGCCCTTT-3′; 5′-TTTCACTCGCCGTTACTAAGG-3′) (Vrain, 1993) using the extracted DNA as a template. The obtained rDNA-ITS sequences were compared to other sequences using BLAST (NCBI), and molecular phylogenetic analysis was constructed using the maximum likelihood method of the MEGA7.0 software, with the number of replicates for the tree-building step set at 1,000 (bootstrap replicates = 1,000). The PCR reaction system was performed as follows: 2 μL DNA template, 2 μL of each primer, 25 μL of 2×Rapid Taq Master Mix, and 21 μL ddH2O. The PCR reaction procedure included initial denaturation at 95 °C for 3 min, followed by 35 cycles of 95 °C for 30 s, 55 °C for 30 s, and 72 °C for 30 s, and a final extension step at 72 °C for 5 min, followed by storage at 4 °C. The PCR products were examined and sequenced. In addition, specific primers were used to detect nematodes: Mp-F/Mp-R (5′-GTTTTCGGCCCGCAACATGT-3′ and 5′-CACCGCCTTGCGTAAACTCC-3′). The PCR reaction system was described above. The PCR reaction procedure included initial denaturation at 95 °C for 3 min followed by 30 cycles of 95 °C for 30 s, 63 °C for 30 s, 72 °C for 45 s, and a final extension at 72 °C for 5 min, followed by storage at 4 °C. Amplification products were examined by 1% agarose gel electrophoresis for the appearance of a single band of the expected size of 409 bp (He et al., 2020).

Healthy Orah seedlings were purchased from the Guangxi Subtropical Crops Research Institute and transplanted into a pot containing sandy loam soil for growth in a greenhouse at room temperature under natural light. After survival, 1,000 J2 nematodes were inoculated into the roots of 10 Orah seedlings, while another 10 Orah seedlings without nematodes were used as a negative control. The root soil was removed after 3 months to observe the damage caused by the nematodes to the roots.

Soil chemical properties

The chemical properties including soil pH, organic matter, total nitrogen (TN), available nitrogen (AN), total phosphorus (TP), available phosphorus (AP), total potassium (TK), and available potassium (AK) were determined as described previously (Bao, 2000). Soil pH was determined using a composite glass electrode meter with a soil: water ratio of 1:2.5. Soil TN was determined using the semi-micro Kjeldahl method; soil AN was determined using a Kjeldahl nitrogen meter; soil TP and AP were determined using the molybdenum-antimony anti-colorimetric method; and soil TK and AK were determined using the flame photometric method.

Soil DNA isolation and PCR conditions

Total DNA was extracted from each rhizosphere soil sample (0.4 g) using the MoBio PowerSoil kit according to the manufacturer’s instructions. The purity and concentration of the extracted DNA were quantified using a Nanodrop spectrophotometer (ND-2000) and on 0.5% agarose gels. Bacterial communities were determined using the universal forward primer (338F:5′-ACTCCTACGGGAGGCAGCAG-3′) and reverse primer (806R:5′-GGACTACHVGGGTWTCTAAT-3′) to amplify the V3–V4 region of the 16S rRNA gene (Guo et al., 2019). Fungal communities were examined using the ITS1 gene, amplified using the universal forward primer (ITS1F:5′- CTTGGTCATTTAGAGGAAGTAA-3′) and reverse primer (ITS2R:5′- GCTGCGTTCTTCATCGATGC-3′) (Li et al., 2020). The PCR reaction was performed in a 20 μL volume containing 5×FastPfu Buffer 4 μL, 2.5 mM dNTPs 2 μL, each primer 0.8 μL, FastPfu Polymerase 0.4 μL, bovine serum albumin (BSA) 0.2 μL, and soil genomic DNA 10 ng. Amplifications were performed using an initial denaturation step of 95 °C for 3 min, followed by 30 cycles of denaturation at 95 °C for 30 s, annealing at 50 °C for 30 s, and extension at 72 °C for 45 s; with a final extension step at 72 °C for 7 min, then stored at 4 °C. The PCR products were purified with a PCR Clean-Up^TM kit (MO BioLabs, Carlsbad, CA, USA) and sent to the Majorbio Company (Shanghai, China) for sequencing using an Illumina MiSeq PE300 (Edgar, 2013).

High-throughput sequencing analysis

The raw sequenced reads were quality-controlled (QC) using fastp software (https://github.com/OpenGene/fastp, version 0.20.0) (Chen et al., 2018), and spliced using FLASH software (http://www.cbcb.umd.edu/software/flash, version 1.2.7) (Magoc & Salzberg, 2011). Quality control involved: filtering bases with quality values below 20 at the end of the reads, setting a window of 50 bp, truncating back-end bases from the window if the average quality value within the window was below 20, filtering reads below 50 bp after QC, and removing reads containing N bases. Pairs of reads were spliced (merged) into one sequence with a minimum overlap length of 10 bp according to the overlap relationship between paired end double-end sequencing reads. The maximum mismatch ratio in the overlap region of the spliced sequence was 0.2, and the non-conforming sequences were screened. The samples were distinguished according to the barcode and primers at the beginning and end of the sequence, and the sequence orientation was adjusted. The number of mismatches in the barcode was 0, and the maximum number of primer mismatches was 2. The data were used for subsequent bioinformatics analyses.

All data analyses were performed using the Meguiar BioCloud platform (https://cloud.majorbio.com). Alpha diversity Coverage, and Chao 1, Simpson, Shannon, and Sobs indices were calculated using mothur software (http://www.mothur.org/wiki/Calculators), and the Wilcoxon rank-sum test was used to analyze group differences in Alpha diversity and to complete the dilution curve analysis; the similarity of microbial community structure between samples was examined using principal coordinate analysis (PCoA) based on the Bray-Curtis distance algorithm; the analysis of microbial community composition was performed with python software (https://www.python.org); the Wilcoxon rank-sum test, two-tailed test, bootstrap algorithm for microbial community inter-group variation and Bray-Curtis distance algorithm for correlation with environmental factors were performed using R-3.3.1 software (R Core Team, 2016).

Statistical analysis

IBM SPSS Statistics 20 was used to analyze significant differences at the p < 0.05 level according to the Duncan multiple range test in the growth indicators (the fruit mass and diameter of Orah), soil chemical properties, and ecological indicators (Coverage, Chao 1, Simpson, and Shannon indices). The redundancy analysis (RDA) through the analysis software R-3.3.1 (vegan) was relied upon to examine the relationships among the soil chemical properties and microbial communities.

Root-knot nematode infection decreased Orah yield

In June, the growth of many Orah infected by root-knot nematode, was weak. Compared with healthy roots, the roots infected by nematodes had root knots of different sizes, and the root disks were together to form fibrous roots, which were messy (Figs. 1A, 1B). The mass and diameter of fruits were significantly lower in root-knot nematode infected Orah than in healthy Orah (Fig. 1C). The average fruit diameter of healthy Orah was 3.86 cm, while the average fruit diameter of root-knot nematode infected Orah was 2.95 cm. The average fruit diameter of root-knot nematode infected Orah was reduced by 23.58% compared to that of healthy Orah (Fig. 1D). The average fruit mass of root-knot nematode infected Orah was 7.49 g, while the average fruit mass of healthy Orah was 19.48 g. The average fruit mass of root-knot nematode infected Orah was reduced by 61.55% compared to that of healthy Orah (Fig. 1E).

Identification of root-knot nematode infecting Orah

There was slight variation among individuals in the female perineal pattern population, but the degree of variation among populations was similar. Microscopic observation of root-knot nematode populations isolated from Orah showed that the root-knot nematode was M. panyuensis: the female was white, spherical to pear-shaped, with an obvious neck, and the neck ring was clear. The back ring was not obvious, and the excretory orifice was located at the median bulb. The pin was developed, and the basal knob was thick. The perineal pattern was oval, the line was smooth, the back arch was low, and there was no obvious side line (Fig. 2A). The morphology of the nematode was consistent with that previously reported (He et al., 2020).

The length of the obtained rDNA-ITS sequence was 869–870 bp (GenBank accession numbers OR135523–OR135524), Blast results confirmed that those sequences were 97.59–99.08% identical to those of M. panyuensis sp. n. from Arachis hypogaea L. in Guangdong, China (AY394719.1) (Liao et al., 2005). The phylogenetic tree was constructed based on the rDNA-ITS sequences, and the results showed that M. panyuensis from Guangxi was clustered with M. panyuensis sp. n. from Guangdong (AY394719.1) (Liao et al., 2005) within a group at a value of 100% (Fig. 2B). A single 409 bp specific fragment was obtained through PCR amplification using the DNA of root-knot nematodes as a template and the specific primers Mp-F/Mp-R of M. panyuensis (Fig. 2C).

Cultured J2s were inoculated into healthy Orah seedlings with good growth and cultured at room temperature for 3 months. The inoculated roots were found to produce root knots (Figs. 2D, 2E).

Soil chemical properties

Infection with M. panyuensis did not significantly affect the pH but had a great influence on organic matter, TN, AN, TP, AP, TK, and AK (Table 1). The organic matter, AP, AK, TN, TP, and TK significantly increased in M. panyuensis-infected Orah rhizosphere soil by 58.87%, 14.39%, 521.39%, 37.14%, 52.31%, and 30.34%, respectively compared with those in healthy Orah rhizosphere soil.

General analyses of the sequencing data

The V3–V4 region of bacterial 16S rDNA and fungal ITS high-throughput sequencing results from Orah rhizosphere soil samples were analyzed, yielding 348,051 and 382,559 optimized sequences, respectively. Furthermore, 3,509 and 1,085 amplicon sequence variant (ASV) sets were generated using the clustering method. The cluster analysis of the ASV sets covered over 99% of the Orah rhizosphere soil microorganisms, indicating that the sequencing results reflected changes in the citrus rhizosphere soil microbial community. The dilution curves of the Shannon and Sobs indices showed a gentle slope with increasing sequencing depth (Fig. S1), suggesting that the sequencing depth was sufficient to capture the vast majority of soil bacteria and fungi. Additionally, the ecological index and coverageindicated that the current sequencing depth was adequate to cover more than 99% of soil bacteria and fungi (Table 2).

Soil bacterial and fungal diversity

The diversity of bacterial and fungal communities in healthy Orah rhizosphere soils was compared with those in M. panyuensis-infected soils. The community richness index (Chao1 index) and community diversity (Shannon index) of bacteria in M. panyuensis-infected Orah soil increased, whereas community diversity (Simpson index) decreased. Meanwhile, the Chao1 and Shannon indices of fungi in M. panyuensis-infected Orah soil decreased, whereas the Simpson index increased. However, there was no significant difference between the healthy and M. panyuensis-infected Orah rhizosphere soils (Table 2). The M. panyuensis-infected Orah rhizosphere soil bacterial community and the healthy one were separated according to PCoA; the combined horizontal and vertical axes in the figure explained 70.6%. The analysis of similarity (ANOSIM) discriminant coefficient R was 0.1852 (Fig. 3A). However, the result was not significantly different (p > 0.05). The M. panyuensis-infected Orah rhizosphere soil fungal community partly overlapped with the healthy Orah soil bacterial community. The combined horizontal and vertical axes explained 68.6% of the variance, and the ANOSIM discriminant coefficient R was 0.3704, but the difference was no significant (p > 0.05, Fig. 3B). These results showed that there was no significant difference in rhizosphere soil community diversity between healthy and M. panyuensis-infected Orah rhizosphere soils.

Soil bacterial and fungal taxonomic composition

The M. panyuensis-infected and healthy Orah rhizosphere soils had the same bacterial community composition, with 229 orders detected. Among them, the relative abundances of Rhizobiales, Burkholderiales, Acidobacteriales, Bacillales, Sphingobacteriales, Bryobacterales, Vicinamibacterales, Frankiales, Chitinophagales, Ktedonobacterales, Gaiellales, and Xanthomonadales were all above 2%, and represented the dominant bacteria in Orah rhizosphere soil. The abundances of the remaining 217 orders were below 2%. The abundance of Burkholderiales in M. panyuensis-infected Orah soil (63%) was significantly higher than the 37% in healthy Orah soil (Fig. 4A). In total, 609 genera were identified. Among them, Bacillus, Bryobacter, Sphingomonas, Acidothermus, Burkholderia-Caballeronia-Paraburkholderia, norank-f-norank-o-Acidobacteriales, norank-f-norank-o-Vicinamibacterales, norank-f-norank-o-Gaiellales and norank-f-Xanthobacteraceae accounted for over 2%, and were the dominant bacteria in Orah rhizosphere soil. The abundances of the other 600 genera were below 2%. The abundance of Burkholderia-Caballeronia-Paraburkholderia in M. panyuensis-infected Orah soil was 84% (Fig. 4B). Which was significantly higher than the 16% in healthy Orah soil. These results indicate that Burkholderia-Caballeronia-Paraburkholderia were closely related to the occurrence of nematode disease.

M. panyuensis-infected and healthy Orah rhizosphere soils had the same fungal community composition, with 11 phyla detected. Among these, the relative abundances of Ascomycota, Basidiomycota, unclassified_k_Fungi, Rozellomycota, and Mortierellomycota were above 1%, and represented the dominant phyla in the rhizosphere soil. The abundances of the other six phyla were below 1%. The abundance of Basidiomycota in M. panyuensis-infected Orah rhizosphere soil was 81%; this was significantly higher than the 19% in healthy Orah rhizosphere soil (Fig. 4C). A total of 275 genera were identified. Among these, Lycoperdon, Roussoella, Fusarium, Neocosmospora, Penicillium, Chaetomium, Talaromyces, Acrocalymma, unclassified_k_Fungi, and Tetragoniomyces accounted for over 3%, whereas the remaining 265 genera accounted for below 3%. The abundance of Lycoperdonin M. panyuensis-infected Orah rhizosphere soil was 99%; this was significantly higher than the 1% in healthy Orah rhizosphere soil (Fig. 4D). This indicated that there was a connection between Lycoperdon and the occurrence of nematode disease.

Comparative analysis between groups of healthy and M. panyuensis-infected Orah soil microbial communities

Comparative analysis of bacteria in Orah rhizosphere soil samples indicated that the relative abundance of Burkholderia-Caballeronia-Paraburkholderia in M. panyuensis-infected Orah soil was 4.71%, which represents a 5.44-fold increase compared to healthy Orah rhizosphere soil groups (0.87%). The relative abundance of Bacillus in M. panyuensis-infected Orah rhizosphere soil was 5.53%, reflecting a 1.40-fold increase over the healthy Orah rhizosphere soil group (3.96%). The relative abundance of Sphingomonas in M. panyuensis-infected Orah rhizosphere soil was 3.75%, representing a 1.50-fold increase compared to healthy Orah rhizosphere soil groups (2.50%) (Fig. 5A). These results further indicated a relationship between Burkholderia-Caballeronia-Paraburkholderia, Bacillus, Sphingomonas, and nematode diseases.

Similarly, the comparative analysis of fungi in Orah rhizosphere soil samples revealed that the relative abundance of Lycoperdon in M. panyuensis-infected Orah rhizosphere soil was 19.55%, a 77.01-fold increase over healthy Orah rhizosphere soil groups (0.2506%). The relative abundance of Fusarium in M. panyuensis-infected Orah rhizosphere soil was 11.06%, which was 1.68-times greater than that in healthy Orah rhizosphere soil groups (6.58%). The relative abundance of Neocosmospora in M. panyuensis-infected Orah soil was 11.22%, indicating a 2.50-fold increase compared to healthy Orah rhizosphere soil groups (4.48%). The relative abundance of Talaromyces in M. panyuensis-infected Orah rhizosphere soil was 7.98%, showing a 12.28-fold increase overthe healthy Orah soil group (0.65%). The relative abundance of Tetragoniomyces in M. panyuensis-infected Orah rhizosphere soil was 3.79%, which represents a 189.50-fold increase compared to the healthy Orah rhizosphere soil group (0.02%) (Fig. 5B). These findings further suggested a relationship between Lycoperdon, Fusarium, Neocosmospora, Talaromyces, Tetragoniomyces, and nematode disease.

Correlation analysis between healthy and M. panyuensis-infected Orah soil microbial communities and environmental factors

RDA showed that soil organic matter and available N, P, and K were significantly and positively correlated with the bacterial community in M. panyuensis-infected Orah rhizosphere soil. The explanatory degrees of the horizontal and vertical coordinates were 46.02% and 28.01%, respectively. Soil organic matter and available K showed strong correlation coefficients (r2) of 0.9982 and 0.9693, respectively (Fig. 6A). Total N, P, and K were significantly positively correlated with the bacterial community in M. panyuensis-infected Orah rhizosphere soil. The interpretations of the horizontal and vertical coordinates were 39.05% and 27.83%, respectively. Total N and K showed a strong positive correlation, with correlation coefficients (r²) of 0.9342 and 0.9055, respectively (Fig. 6B). In addition, four of the ten genera with the highest relative abundance were positively correlated with the bacterial community in M. panyuensis infected Orah rhizosphere soil, including Burkholderia-Caballeronia-Paraburkholderia (Figs. 6A, 6B). These results add further evidence that Burkholderia-Caballeronia-Paraburkholderia were related to nematode infection.

The soil organic matter and available N, P, and K were significantly positively correlated with the fungal community in M. panyuensis-infected Orah rhizosphere soil. The horizontal and vertical coordinates were 58.18% and 20.09%, respectively. Soil organic matter showed a strong positive correlation with a correlation coefficient (r²) of 0.9116 (Fig. 6C). Total N, P, and K were significantly positively correlated with the fungal community in M. panyuensis-infected Orah rhizosphere soil. The interpretations of the horizontal and vertical coordinates were 47.58% and 20.74%, respectively. Among these, total P and total K showed strong positive correlations, with correlation coefficients (r²) of 0.9031 and 0.8856, respectively (Fig. 6D). In addition, four of the ten genera with the highest relative abundance were positively correlated with the fungal community in the M. panyuensis-infected Orah rhizosphere soil, including Lycoperdon (Figs. 6C, 6D). These results provided further evidence that Lycoperdon was related to nematode infection.