First report of rice root-knot nematode, Meloidogyne graminicola, infecting Juncus microcephalus in Brazil

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Journal of Nematology

Society of Nematologists

Subject: Life Sciences

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ISSN: 0022-300X
eISSN: 2640-396X

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First report of rice root-knot nematode, Meloidogyne graminicola, infecting Juncus microcephalus in Brazil

Cristiano Bellé * / Paulo Sergio dos Santos / Tiago Edu Kaspary

Keywords : Identification, Molecular analyses, EST phenotype, Root-knot nematodes

Citation Information : Journal of Nematology. Volume 53, Pages 1-4, DOI: https://doi.org/10.21307/jofnem-2021-031

License : (CC-BY-4.0)

Received Date : 29-January-2021 / Published Online: 20-March-2021

ARTICLE

ABSTRACT

Juncus microcephalus plants showing symptoms of root-knot nematode infestation were observed in the municipality of Agudo, Rio Grande do Sul state, Brazil. Based on morphological observation, esterase phenotypes, and molecular analyses of rDNA-ITS and D2-D3 regions of 28S rDNA, the causal agent of the observed symptoms was identified as Meloidogyne graminicola. Pathogenicity of M. graminicola was confirmed by fulfilling modified Koch’s postulates. To our knowledge, this is the first report of M. graminicola in J. microcephalus in Rio Grande do Sul State, Brazil.

Graphical ABSTRACT

Weeds infest agricultural fields in all productive regions of the world. The presence of these species, infesting commercial crops, reduce potential of productivity by competing for environmental resources, allelopathic effects, and serving as alternative hosts for pests and pathogens (Bellé et al., 2019; Webster and Nichols, 2012). The Juncus microcephalus (South American rush) is a weed species originally from south America and is present in wetlands cultivated in many countries worldwide (Balslev and Stefano, 2015; Pivari et al., 2019; Rolon et al., 2010).

Juncus microcephalus Kunth (family Juncaceae) is a persistent and herbaceous weed which develops in flooded areas. This weed blooms and fructifies irregularly throughout the year, with greater seed production in the summer (Balslev and Stefano, 2015). South American rush is present in rice fields as weed throughout the development of rice, competing with the crop and reducing its productive potential. In addition, this weed develops during the absence of crop, serving as an alternative host for many crop pests including arthropods, pathogens, including nematodes.

In October 2020, samples of South American reeds showing many galls on the roots (Fig. 1A, B) were collected in rice fields, before sowing the crop from the municipality of Agudo (29° 34′16, 7″S; 53° 17′17, 4″ O; 53 m), state of Rio Grande do Sul, Brazil. No symptoms were observed in the aerial part of infected South American rush plants.

Figure 1:

Meloidogyne graminicola Golden and Birchfield, 1965, root infestation symptoms on South American rush (Juncus microcephalus Kunth). Root-knot symptoms of galls of J. microcephalus from the field (A, B) and in the greenhouse (C, D).

10.21307_jofnem-2021-031-f001.jpg

This species was identified from esterase using esterase phenotypes (n = 20 females) (Carneiro and Almeida 2001; Carneiro et al., 2000), morphological measurement of second-stage juveniles (J2) (n = 20), females (n = 10) and males (n = 10), and perineal patterns (n = 20) and through the amplification and sequencing of ITS1-5.8S-ITS2 rRNA region and the D2 to D3 fragment of the 28S ribosomal RNA gene (De Ley et al., 1999; Schmitz et al., 1998). Genomic deoxyribonucleid acid (DNA) was ultimately obtained from females using the NaOH method (Stanton et al., 1998).

The nematode population density observed in the sample was 1,980 J2/g of J. microcephalus roots. The J2s had the following morphometric characters: length (L) = 499.5 ± 45.0 (389.0-500.5) μm, a = 26.5 ± 1.1 (24.5-30.0), c = 5.1 ± 0.4 (4.90-7.1), stylet length =14.9 ± 0.5 (13.3-16.1) μm, dorsal esophageal gland opening (DGO) = 3.6 ± 0.4 (3.1-4.5) μm, tail length = 70.2 ± 3.2 (61.5-79.9) μm and hyaline tail terminus = 18.0 ± 1.3 (15.1-23.5) μm. Morphological measurements of females included L = 690.5 ± 30.5 (500.5-799.5) μm, stylet length = 13.5 ± 0.3 (11.2-14.7) μm, and DGO = 3.6 ± 0.4 (3.1-5.5) μm. The female’s perineal patterns were oval shape and a low dorsal arch without the presence of a lateral lines and the cuticular striations were smooth and thick in the dorsal region of the vulva. Male measurements were L = 1,401.5 ± 150.5 (1,150.5-1,792.0) μm, stylet length = 19.5 ± 0.5 (17.9-20.2) μm, DGO = 3.5 ± 0.5 (2.4-4.0) μm, tail = 11.4 ± 1.5 (9.4-14.0) μm, spicule = 31.0 ± 1.4 (28.0-36.5) μm. The overall morphology and morphometrics of the population of fit into Meloidogyne graminicola (Golden and Birchfield, 1965) according to the original description (Golden and Birchfield, 1965).

The polymorphisms of the esterase bands observed by electrophoresis revealed the phenotype VS-1 (G1) (Rm = 0.70) typical of M. graminicola (Carneiro et al., 1996). The sequences of the rDNA regions (ITS: 433 bp and D2-D3 of 28S: 446 bp) were submitted to GenBank (ITS: MW537706 and D2-D3 of 28S: MW537709). Searches on BLAST showed 99 to 100% identity with sequences of M. graminicola isolates from Brazil, Taiwan, and China.

To satisfy a modified Koch’s postulates, J. microcephalus plantlets were grown in 1.7 L pots filled with a sterilized soil. The seeds were obtained directly from the weeds in the rice fields. Seeds were sown in trays filled with commercial substrate. In total, 20 days after emergence, the seedlings were transplanted to pots (one per pot), five days after transplanting, six plantlets were inoculated with 2,000 eggs and J2s from the original population of M. graminicola, extracted with 0.5% NaOCl according to Hussey and Barker (1973), using a blender instead of manual shaking. In addition, non-inoculated control six plants were also included in the study. Plants were maintained under greenhouse conditions at 25 ± 3°C, with watering as needed. After 60 days, the inoculated plants exhibited galled root systems similar observed in the field, with a nematode reproduction factor (final population/initial population) of 9.5 (Fig. 1C, D). Plants did not exhibit any above-ground symptoms. The non-inoculated plants did not exhibit any galls.

Rice plants (cultivars IRGA 424 RI, Guri Inta CL, IRGA 431 CL, and Puita Inta CL) five days after emergence, seedlings were transplanted to 1.7 L pots containing sterilized soil, one plant per pot. Plants were inoculated five days after transplanting were also inoculated with an isolate of M. graminicola from J. microcephalus using the same methods as described above. The plants exhibited galled root systems with a nematode reproduction factor ranging from 12.4 to 25.5 (Fig. 2A-D). Inoculated plants showed a reduction in fresh weight of shoots (25-29%) and roots (27-30%) compared to non-inoculated plants. These results confirmed the pathogenicity of the M. graminicola in J. microcephalus and rice.

Figure 2:

Pathogenicity of Meloidogyne graminicola Golden and Birchfield, 1965, isolated from Juncus microcephalus Kunth, in different rice cultivars (A: IRGA 424 RI, B: Guri Inta CL, C: IRGA 431 CL and D: Puita Inta CL).

10.21307_jofnem-2021-031-f002.jpg

An effective control of nematodes, limiting the increase in the population of these phytoparasites is the proper management of weeds throughout the year including the period of absence of crops where only weeds serves the hosts for this phytoparasite (Bellé et al., 2016). In this sense, the control method most used and considered most efficient for the management of J. microcephalus is the chemical with the use of herbicides. In this way, by controlling the weeds, there will be no shelter and food for the nematodes, leading to gradual reduction in their population.

Therefore, the management of weeds is of great importance, since it directly impacts the survival of Meloidogyne spp. in agricultural fields, while the negative potential of weeds species on commercial crops is exacerbated when they become phytoparasite hopers. However, based on the knowledge of M. graminicola polyphagia and its host range, effective strategies can be devised in the management of this pathogen, reducing the damage caused to commercial crops. Finally, the reduction of agricultural losses caused by weeds and nematodes can be minimized with the integrated management of these two problems, which are interconnected and enhanced when they occur concurrently. To our knowledge, this is the first report of M. graminicola parasitizing J. microcephalus in Brazil and elsewhere.

References


  1. Balslev, H. and Stefano, D. F. 2015. La familia Juncaceae en México. Acta Botanica Mexicana 111:61–164.
  2. Bellé, C. , Kaspary, T. E. , Schmitt, J. and Kuhn, P. R. 2016. Meloidogyne ethiopica and Meloidogyne arenaria parasitizing Oxalis corniculata in Brazil. Australasian Plant Disease Notes 11:24–27.
  3. Bellé, C. , Kaspary, T. E. , Balardin, R. R. and Antoniolli, Z. I. 2019. Detection of Meloidogyne graminicola parasitising Cyperus rotundus in Rio Grande do Sul, Brazil. Australasian Plant Disease Notes 14:2.
  4. Carneiro, R. M. D. G. and Almeida, M. R. A. 2001. Técnica de eletroforese usada no estudo de enzimas dos nematoides de galhas para identificação de espécies. Nematologia Brasileira 25:555–560.
  5. Carneiro, R. M. D. G. , Almeida, M. R. A. and Carneiro, R. G. 1996. Enzyme phenotypes of Brazilian populations of Meloidogyne spp. Fundamental & Applied Nematology 19:555–560.
  6. Carneiro, R. M. D. G. , Almeida, M. R. A. and Quénéhervé, P. 2000. Enzyme phenotypes of Meloidogyne spp. populations. Nematology 2:645–654.
  7. De Ley, P. , Félix, M. A. , Frisse, L. M. , Nadler, S. A. , Sternberg, P. W. and Thomas, W. K. 1999. Molecular and morphological characterization of two reproductively isolated species with mirror-image anatomy (Nematoda: Cephalobidae). Nematology 1:519–612.
  8. Golden, A. M. and Birchfield, W. 1965. Meloidogyne graminicola (Heteroderidae) a new species of root-knot nematode from grass. Proceedings of the Helminthological Society of Washington 32:228–231.
  9. Hussey, R. S. and Barker, K. R. 1973. A comparison of methods colleting inocula of Meloidogyne spp. including a new technique. Plant Disease Reporter 57:1025–1028.
  10. Pivari, M. O. D. , Melo, P. H. A. , Souza, F. S. , Stehmann, J. R. , Moura-Júnior, E. G. , Moreira, S. N. , Pott, V. J. , Pott, A. , Lopes, A. , Moço, M. C. C. , Oliveira, L. S. , Lins, A. L. A. , Arruda, R. , Morais, I. L. , Silva, G. S. and Ferreira, R. M. 2019. New initiatives for Brazilian aquatic plant data management. Acta Botanica Brasilica 33:78–87.
  11. Rolon, A. S. , Homem, H. F. and Maltchik, L. 2010. Aquatic macrophytes in natural and managed wetlands of Rio Grande do Sul State, Southern Brazil. Acta Limnologica Brasiliensia 22:133–146.
  12. Schmitz, B. , Bürgermeister, W. and Braasch, H. 1998. Molecular genetic classification of central European Meloidogyne chitwoodi and M. fallax populations. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes 50:310–317.
  13. Stanton, J. M. , McNicol, C. D. and Steele, V. 1998. Non-manual lysis of second-stage Meloidogyne juveniles for identification of pure and mixed samples based on the polymerase chain reaction. Australasian Plant Pathology 27:112–115.
  14. Webster, T. M. and Nichols, R. L. 2012. Changes in the prevalence of weed species in the major agronomic crops of the Southern United States: 1994/1995 to 2008/2009. Weed Science 60:145–157.
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FIGURES & TABLES

Figure 1:

Meloidogyne graminicola Golden and Birchfield, 1965, root infestation symptoms on South American rush (Juncus microcephalus Kunth). Root-knot symptoms of galls of J. microcephalus from the field (A, B) and in the greenhouse (C, D).

Full Size   |   Slide (.pptx)

Figure 2:

Pathogenicity of Meloidogyne graminicola Golden and Birchfield, 1965, isolated from Juncus microcephalus Kunth, in different rice cultivars (A: IRGA 424 RI, B: Guri Inta CL, C: IRGA 431 CL and D: Puita Inta CL).

Full Size   |   Slide (.pptx)

REFERENCES

  1. Balslev, H. and Stefano, D. F. 2015. La familia Juncaceae en México. Acta Botanica Mexicana 111:61–164.
  2. Bellé, C. , Kaspary, T. E. , Schmitt, J. and Kuhn, P. R. 2016. Meloidogyne ethiopica and Meloidogyne arenaria parasitizing Oxalis corniculata in Brazil. Australasian Plant Disease Notes 11:24–27.
  3. Bellé, C. , Kaspary, T. E. , Balardin, R. R. and Antoniolli, Z. I. 2019. Detection of Meloidogyne graminicola parasitising Cyperus rotundus in Rio Grande do Sul, Brazil. Australasian Plant Disease Notes 14:2.
  4. Carneiro, R. M. D. G. and Almeida, M. R. A. 2001. Técnica de eletroforese usada no estudo de enzimas dos nematoides de galhas para identificação de espécies. Nematologia Brasileira 25:555–560.
  5. Carneiro, R. M. D. G. , Almeida, M. R. A. and Carneiro, R. G. 1996. Enzyme phenotypes of Brazilian populations of Meloidogyne spp. Fundamental & Applied Nematology 19:555–560.
  6. Carneiro, R. M. D. G. , Almeida, M. R. A. and Quénéhervé, P. 2000. Enzyme phenotypes of Meloidogyne spp. populations. Nematology 2:645–654.
  7. De Ley, P. , Félix, M. A. , Frisse, L. M. , Nadler, S. A. , Sternberg, P. W. and Thomas, W. K. 1999. Molecular and morphological characterization of two reproductively isolated species with mirror-image anatomy (Nematoda: Cephalobidae). Nematology 1:519–612.
  8. Golden, A. M. and Birchfield, W. 1965. Meloidogyne graminicola (Heteroderidae) a new species of root-knot nematode from grass. Proceedings of the Helminthological Society of Washington 32:228–231.
  9. Hussey, R. S. and Barker, K. R. 1973. A comparison of methods colleting inocula of Meloidogyne spp. including a new technique. Plant Disease Reporter 57:1025–1028.
  10. Pivari, M. O. D. , Melo, P. H. A. , Souza, F. S. , Stehmann, J. R. , Moura-Júnior, E. G. , Moreira, S. N. , Pott, V. J. , Pott, A. , Lopes, A. , Moço, M. C. C. , Oliveira, L. S. , Lins, A. L. A. , Arruda, R. , Morais, I. L. , Silva, G. S. and Ferreira, R. M. 2019. New initiatives for Brazilian aquatic plant data management. Acta Botanica Brasilica 33:78–87.
  11. Rolon, A. S. , Homem, H. F. and Maltchik, L. 2010. Aquatic macrophytes in natural and managed wetlands of Rio Grande do Sul State, Southern Brazil. Acta Limnologica Brasiliensia 22:133–146.
  12. Schmitz, B. , Bürgermeister, W. and Braasch, H. 1998. Molecular genetic classification of central European Meloidogyne chitwoodi and M. fallax populations. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes 50:310–317.
  13. Stanton, J. M. , McNicol, C. D. and Steele, V. 1998. Non-manual lysis of second-stage Meloidogyne juveniles for identification of pure and mixed samples based on the polymerase chain reaction. Australasian Plant Pathology 27:112–115.
  14. Webster, T. M. and Nichols, R. L. 2012. Changes in the prevalence of weed species in the major agronomic crops of the Southern United States: 1994/1995 to 2008/2009. Weed Science 60:145–157.

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