Host status of morning-glory (Ipomoea spp.) to Meloidogyne species

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Host status of morning-glory (Ipomoea spp.) to Meloidogyne species

Tiago Edu Kaspary / Ismail Teodoro de Souza Júnior / Rodrigo Ferraz Ramos / Cristiano Bellé *

Keywords : Weeds, Root-knot nematode, Ipomoea spp., Reproduction, Susceptibility

Citation Information : Journal of Nematology. Volume 53, Pages 1-6, DOI: https://doi.org/10.21307/jofnem-2021-018

License : (CC-BY-4.0)

Received Date : 09-September-2020 / Published Online: 15-February-2021

ARTICLE

ABSTRACT

Weeds can be hosting and alternative multipliers of root-knot nematodes (Meloidogyne spp.). Among the main weeds, species of the genus Ipomoea stands out for their cosmopolitan presence and the negative impact on crops. In addition, they can behave as hosts and promote the reproduction of pests, diseases, and nematodes. However, the ability of Meloidogyne nematodes to infect morning-glory (Ipomoea spp.) is little understood. In this context, the objective was to evaluate the reproduction of M. arenaria, M. enterolobii, M. ethiopica, M. hapla, M. incognita, M. javanica, M. luci, and M. morocciensis in I. grandifolia, I. hederifolia, I. nil, I. purpurea, and I. quamoclit. Plants were individually inoculated with 5,000 eggs and second-stage juveniles and kept in a greenhouse for 60 days. The design was completely randomized with six repetitions. After this period, the root system of each plant was evaluated to gall index (IG) and reproduction factor (RF). It was verified that the eight species of Melodoigyne have the capacity to parasitize I. grandifolia, I. hederifolia, I. nil, I. purpurea, and I. quamoclit, showing the susceptibility of these weeds to the plant-parasitic nematodes. The highest RF were observed for M. enterolobii with values of 12.5 and 12.9 for I. quamoclit and I. hederifolia, respectively. While M. arenaria obtained the lowest values, with RF ≤ 4.0 for all species of Ipomoea. Thus, weed species of the Ipomoea genus are potential hosts and multipliers of root-knot nematodes, making it important to be considered in integrated management strategies for these plant-parasitic nematodes.

Graphical ABSTRACT

Weeds are an important biotic factor that limits agricultural crops. The damage caused to the development and productivity of crops can be generated by direct competition for resources, such as water, light, and nutrients, or by the action of allelochemicals produced by weeds (Ramesh et al., 2017). In addition, the presence of weeds can compromise the final quality of agricultural products, by contamination with their vegetable remains and seeds, or by becoming alternative hosts of pests and diseases, multiplying these and becoming a source of inoculum for future infestations (Anwar et al., 2009; Bellé et al., 2017b; Singh et al., 2010).

Currently, among the most problematic weeds for agriculture, the genus Ipomoea stands out, popularly known as morning-glory. Among the main species of this genus, some are of agricultural importance in annual and perennial crops and are reported as weeds in 40 crops in more than 40 countries (Holm et al., 1997; Chauhan and Abugho, 2012), especially Ipomoea hederifolia L., Ipomoea quamoclit L., Ipomoea purpurea L., Ipomoea grandifolia (Dammer) O’Donell and Ipomoea nil (L.) Roth (Kissmann and Groth, 1999). The main characteristic of these plants are the volatile stems and branches, which give rise to the climbing habit (Lorenzi, 2000). In addition to competing with culture, these plants can interfere with cultural practices, especially mechanized harvesting, whose operational efficiency of the harvester is reduced by the fact that the plants are wrapped around the culms of the crop (Elmore et al., 1990; Lorenzi, 2000). These factors, combined with a wide geographical distribution, high production capacity and propagation of propagules and the difficulty in handling amplify the negative effect of weeds on agriculture.

The knowledge of weeds that act as alternative hosts for pests and diseases have been used as an integrated management tool in several agricultural crops. In this context, several weed species have been reported as hosts of plant-parasitic nematodes, among which the root-knot nematodes (Meloidogyne spp. Göldi). This phytoparasite genus has the greatest impact on crops in the world, in addition to being the genus most frequently found in parasitic weed roots (Bellé et al., 2019; Ferraz et al., 1978; Moens and Perry, 2009). In Brazil, an increasing number of studies have been developed that report weeds, present in agricultural areas, as natural hosts of several species of nematodes of the genus Meloidogyne (Bellé et al., 2016; Groth et al., 2017; Kaspary et al., 2017).

Management options for root-knot nematodes are limited, due to their wide range of hosts across various cultures and the lack of effective and environmentally acceptable control methods (McSorley, 1992; Ritzinger and Fancelli, 2006; Starr et al., 2007). There are few cultivars with resistance to root-knot nematodes, and the existing ones are restricted to one or two species of this nematode (Sikora et al., 2018). Due to plant and nematicide resistance limitations, crop rotation is the nematode control method widely used by producers in different parts of the world (Bellé et al., 2020; Ramos et al., 2019). However, the effectiveness of crop rotation for the management of root-knot nematodes is drastically reduced due to the presence of weeds in crops and during the off-season (Bellé et al., 2016; Koenning et al., 1999).

The root-knot nematodes M. javanica (Treub) Chitwood, M. incognita (Kofoid and White) Chitwood, M. arenaria (Neal) Chitwood and M. enterolobii Uang and Eisenback are distributed in the main areas of agricultural production in Brazil. M. hapla Chitwood, M. morocciensis Rammah and Hirschmann. M. luci Carneiro et al. and M. ethiopica Whitehead, on the other hand, are milder climate species, being more limited to these regions. Some of these are of special interest (M. enterolobii and M. ethiopica) due to their ability to overcome some root-knot nematode resistance genes (Carneiro et al., 2016; Galbieri et al., 2020). Currently, little is known about the reproduction of root-knot nematodes in species of morning-glory (Ipomoea spp.) in Brazil. In this context, this study aimed to evaluate the reproduction of eight species of Meloidogyne in Ipomoea grandifolia, I. hederifolia, I. nil, I. purpurea, I. quamoclit.

The experiment was conducted a greenhouse, in independent trials from November 2018 and February 2019, where the reaction of five species of morning-glory (I. grandifolia, I. hederifolia, I. nil, I. purpurea, and I. quamoclit) to eight Meloidogyne species, in a greenhouse a 25°C±3°C. Weeds were identified and classified according to Ferreira and Miotto (2009). A completely randomized experimental design was used with six replications.

It was used with a substrate for the experiment a mixture of soil (Rhodic Ferralsol – WRB 2015) and sand, at proportion of 1:2, with autoclave disinfection before use. The soil was composed of clay=48%; water pH value=6.4; Shoemaker MacLean, and Pratt (SMP) test=6.6; organic matter=3.3%; P=10.7 mg dm−3; K=86 mg dm−3; Ca=5.2 cmolc dm−3; Mg=4.9 cmolc dm−3; and S=8.5 cmolc dm−3. In total, 15 days after emergence of morning-glory on plant were transplanted in each put containing 2,000 dm−3 the substrate.

As inoculum of the root-knot nematode, pure populations of M. arenaria (Est A2), M. enterolobii (Est M2), M. ethiopica (Est E3), M. hapla (Est H1), M. incognita (Est I2), M. javanica (Est J3), M. luci (Est L3), and M. morocciensis (Est A3) confirmed using the electrophoresis technique for isoenzymes, as proposed by Carneiro and Almeida (2001), were maintained and multiplied in ‘Santa Cruz’ tomato plants (Solanum licopersycum L.). The nematode inoculum was obtained from the root system of tomato plants kept in a greenhouse, using the method of Hussey and Barker (1973), extracted with 0.5% NaOCl, using a blender instead of manual shaking. The experimented plants were inoculated five days after transplantation with 3 ml. of suspension containing 5,000 eggs and second juvenile (J2) per plant in three holes approximately two cm deep around the plant. ‘Santa Cruz’ tomatoes were used as a control of the viability of the inoculum of the nematode population.

The roots of each morning-glory plant were washed individually 60 days after inoculation. The galls were counted following the methodology proposed by Taylor and Sasser (1978), where 0=no galls, 1=1 to 2, 2=3 to 10, 3=11 to 30, 4=31 to 100, and 5=more than 100 galls per root system. Subsequently, the root systems were processed following the method of Hussey and Barker (1973), to obtain the final number of nematodes. From this, the reproduction factor (final population/initial population) was determined, as described by Oostenbrink (1966). Thus, each morning-glory species was classified as RF=0 for immune, RF <1 for resistant, and RF>1 was susceptible for each nematode species.

The similarity form, the experiment was performed twice times and compared by preliminary analysis of variance (ANOVA). This analysis was not significant for first and second round, allowing the use of the data together. Subsequently, the treatments were compared by Scott-Knott clustering test at probability level of 5%, using the GENES software (Cruz, 2006).

The results obtained demonstrate a similar behavior of the Ipomoea species evaluated in relation to the reaction to the different Meloidogyne species. The viability of the inoculum was confirmed by the reproduction factor above 17.5 found in S. lycopersicum, used as a control, for all nematode species evaluated (Table 1). The gall indexes presented by the five Ipomoea species were greater than 4.5 when they were subjected to the eight species of root-knot nematode (Table 1).

Table 1.

Galls index (GI), and factor of reproduction (RF) of M. arenaria, M. enterolobii, M. ethiopica, M. hapla, M. incognita, M. javanica, M. luci, and M. morocciensis in I. grandifolia, I. hederifolia, I. nil, I. purpurea, and I. quamoclit.

10.21307_jofnem-2021-018-t001.jpg

The species I. grandifolia and I. purpurea presented the largest reproduction factors for M. javanica, with an RF of 9.7 and 8.9, respectively (Table 1). While for M. incognita, the highest index was observed in I. hederifolia (RF=9.0), whereas I. nil obtained the lowest reproduction factor for this nematode (RF=5.7). M. arenaria showed greater capacity to multiply in plants of the species I. quamoclit, I. purpurea and I. hederifolia with RFs greater than 2.7 on I. grandifolia (Table 1). Plants of the species I. grandifolia, I. nil, and I. purpurea were more likely to multiply the nematode M. hapla when compared to I. hederifolia (Table 1). For M. ethiopica, the RFs varied from 7.1 to 10.1, with the weeds I. hederifolia, I. quamoclit, and I. grandifolia having the highest RF (Table 1). Whereas the species I. quamoclit and I. hederifolia also resulted in the highest RF for the nematode M. enterolobii, with reproduction factors of 12.5 and 12.9, respectively (Table 1). For M. luci, the species I. nil, I. hederifolia and I. grandifolia presented the highest RF, varying from 8.5 to 9.3, while for I. quamoclit, the lowest reproduction factor was observed for this nematode (RF=3.6).

Species of Ipomea showed little variation in the reproductive capacity of M. morocciensis, however, I. hederifolia, I. grandifolia and I. purpurea showed the highest RF statistically, ranging from 5.5 to 6.5 (Table 1). In contrast, I. nil, was the species that presented the lowest RF values for most (6) of the Meloidogyne species evaluated, while I. hederifolia was the one that presented the highest RF values for six of the eight Meloidogyne species evaluated.

From the RF averages of the different plant-parasitic nematodes facing the different species of Ipomoea, it was found that M. enterolobii presenting the highest reproduction capacity with RF of 11.32 (Fig. 1). This value is about 70 and 62% higher than values found for M. arenaria and M. hapla, with an average RF of 3.44 and 4.32, respectively. For the other species, the mean RF varied from 4.42 to 8.62, values considered high, demonstrating to which species of the genus Ipomea are excellent multipliers of the root-knot nematodes (Fig. 1).

Figure 1:

Average reproduction factor of Meloidogyne species, multiplied in I. grandifolia, I. hederifolia, I. nil, I. purpurea, and I. quamoclit. *Means followed by the same letter in the columns do not differ by the Scott-Knott test at 5% probability.

10.21307_jofnem-2021-018-f001.jpg

The high capacity to host and multiply the root-knot nematode species by the five Ipomoea species in the present study indicates that these weeds behaved as susceptible to these phytoparasites, the presence of which is the guarantee of the perpetuation of the plant-parasitic nematodes in cultivation areas (Bellé et al., 2017b; Singh et al., 2013). In addition, species of the genus Ipomoea have different germination flows as an aggravating factor throughout the year depending on temperature and humidity (Azania et al., 2009; Carvalho et al., 2008). This factor enhances the maintenance and reproduction of populations of different root-knot nematodes during the year.

The polyphagous capacity presented by the root-knot nematodes, care must be taken with the choice of the cultures to be implanted and the species in succession, whether commercial culture or not, in order to minimize the damage caused by M. arenaria, M. enterolobii, M. ethiopica, M. hapla, M. incognita, M. javanica, M. luci and M. morocciensis. Even in this context, weed control is of even greater importance, due to the proven and high susceptibility and potential host to the nematode species under study, demonstrating the polyphagous action of the pathogen, which can have negative consequences for the cultivation areas where they are located, mainly in areas with soybeans, beans, sugarcane, tobacco, vegetables, and fruit, where plant-parasitic nematodes are already widespread (Bellé et al., 2017b).

Considering the management of plant-parasitic nematodes, a way to control population growth population during the period of presence of cultivation of economic interest, is the proper management of weeds throughout the year (Bellé et al., 2016). The control method most used and considered most efficient for the management of morning-glory is the chemical with the use of pre and post-emergence herbicides, getting satisfactory control (Azania et al., 2009). In post-emergence, the use of herbicides is recommended only in the early stage of the morning-glory (2-4 leaves). Although to date, there are no reports of Ipomoea spp. resistant to different herbicides (Heap, 2020).

Thus, the difficulty in controlling weeds increases the management complexity of Meloidogyne spp. aggravating the harmful potential of the presence of these plants in crop fields. However, from the knowledge of the host range of M. javanica, M. incognita, M. arenaria, M. hapla, M. ethiopica, M. enterolobii, M. luci, and M. morocciensis, it will be possible to trace effective strategies in the management of these pathogens, reducing the damage caused to commercial crops. Finally, the reduction of agricultural losses caused by weeds and phytonematodes can be minimized with the integrated management of these two problems, which are interconnected and enhanced when they occur concurrently.

References


  1. Anwar, S. S., Zia, A. and Javed, N. 2009. Meloidogyne incognita infection of five weed genotypes. Pakistan Journal of Zoology 41:95–100.
  2. Azania, C. A. M. , Marques, R. P. , Azania, A. A. P. M. and Rolim, J. C. 2009. Overcoming dormancy of Ipomoea quamoclit and I. hederifolia seeds. Planta Daninha 27:23–27.
  3. 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.
  4. Bellé, C. , Kulczynski, S. M. , Kaspary, T. E. and Kuhn, P. R. 2017b. Weeds as host alternative to Meloidogyne incognita. Nematropica 47:26–33.
  5. Bellé, C. , Kaspary, T. E. , Balardin, R. R. , Ramos, R. F. and Antoniolli, Z. I. 2019. Meloidogyne species associated with weeds in Rio Grande do Sul. Planta Daninha 37:e019214250.
  6. Bellé, C. , Ramos, R. F. , Balardin, R. R. , Nora, D. D. and Kaspary, T. E. 2020. Multiplication potential of Meloidogyne arenaria in weeds found in Brazil. European Journal of Plant Pathology 157:441–447.
  7. 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.
  8. Carneiro, R. M. D. G. , Monteiro, J. M. S. , Silva, U. C. and Gomes, G. 2016. “Gênero Meloidogyne: diagnose através de eletroforese de isoenzimas e marcadores SCAR”, In Oliveira, C. M. , Santos, M. A. and Castro, L. H. S. (Eds), Diagnose de fitonematoides Millennium, Campinas, pp. 71–93.
  9. Carvalho, S. J. P., López-Ovejero, R. F. and Christoffoleti, P. J. 2008. Growth and development of five weed species of the Amaranthus genus. Bragantia 67:317–.
  10. Chauhan, B. S. and Abugho, S. B. 2012. Threelobe morningglory (Ipomoea triloba) germination and response to herbicides. Weed Science 60:199–204.
  11. Cruz, C. D. 2006. Programa GENES: biometria UFV, Vicosa.
  12. Elmore, C. D. , Hurst, H. R. and Austin, D. F. 1990. Biology and control of morning-glories (Ipomoea spp.). Weed Science 5:83–114.
  13. Ferraz, L. C. C. B. , Pitelli, R. A. and Furlan, V. 1978. Nematoides associados a plantas daninhas na região de Jaboticabal, SP-primeiro relato. Planta Daninha 1:5–11.
  14. Ferreira, P. P. A. and Miotto, S. T. S. 2009. Synopsis of Ipomoea L. (Convolvulaceae) species in Rio Grande do Sul State, Brazil. Revista Brasileira de Biociências 7:440–453.
  15. Galbieri, R. , Davis, R. F. , Scoz, L. B. , Belot, J. L. and Skantar, A. M. 2020. First Report of Meloidogyne enterolobii on Cotton in Brazil. Plant Disease 104:2295, available at: https://doi.org/10.1094/PDIS-02-20-0365-PDN.
  16. Groth, M. Z , Bellé, C. , Cocco, K. L. T. , Kaspary, T. E. , Casarotto, G. , Cutti, L. and Schmitt, J. 2017. First report of Meloidogyne enterolobii infecting the weed Jerusalem cherry (Solanum pseudocapsicum) in Brazil. Plant Disease 101:510.
  17. Heap, I. 2020. The International Survey of Herbicide Resistant Weeds, available at: http://www.weedscience.org.
  18. Holm, L. , Doll, J. , Holm, E. , Pancho, J. and Herberger, J. 1997. World weeds: natural histories and distribution Wiley, New York.
  19. 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.
  20. Kaspary, T. E. , Bellé, C. , Groth, M. Z. , Cocco, K. L. T. , Cutti, L. and Casarotto, G. 2017. Amaranthus viridis is a weed host of Meloidogyne arenaria in Rio Grande do Sul State, Brazil. Plant Disease 101:635.
  21. Kissmann, K. G. and Groth, D. 1999. “Convolvulaceae Juss, Vol. 1”, In Kissmann, K. G. and Groth, D. (Eds), Plantas infestantes e nocivas 2nd ed., BASF, São Paulo, pp. 673–693.
  22. Koenning, S. R. , Overstreet, C. , Noling, J. W. , Donald, P. A. , Becker, J. O. and Fortnum, B. A. 1999. Survey of crop losses in response to phytoparasitic nematodes in the United States for 1994. Journal of Nematology 31:587–618.
  23. Lorenzi, H. 2000. Plantas daninhas do Brasil: terrestres, aquáticas, parasitas e tóxicas Instituto Plantarum, Nova Odessa.
  24. McSorley, R. 1992. Nematological problems in tropical and subtropical fruit tree crops. Nematropica 22:103–116.
  25. Moens, M. and Perry, R. 2009. Migratory plant endoparasitic nematodes: a group rich in contrasts and divergence. Annual Review Phytopathology 37: 313–32.
  26. Oostenbrink, M. 1966. Major characteristics of the relation between nematodes and plants. Mendelingen Landbouwhogeschool, Wageningen 66:1–46.
  27. Ramesh, K. , Matloob, A. , Aslam, F. , Florentine, S. K. and Chauhan, B. S. 2017. Weeds in a changing climate: vulnerabilities, consequences, and implications for future weed management. Frontiers in Plant Science 8:95.
  28. Ramos, R. F. , Kaspary, T. E. , Balardin, R. R. , Nora, D. D. , Antonioli, Z. I. and Bellé, C. 2019. Plantas daninhas como hospedeiras dos nematoides-das-galhas. Revista Agronomia Brasileira 3:rab201906.
  29. Ritzinger, C. H. S. P. and Fancelli, M. 2006. Integrated management of nematodes in the banana tree culture. Revista Brasileira de Fruticultura 28:331–338.
  30. Sikora, R. , Coyne, D. , Hallmann, J. and Timper, P. 2018. Plant parasitic nematodes in subtropical and tropical agriculture CAB International, Wallingford.
  31. Singh, S. K. , Ash, G. J. and Banks, N. C. 2013. Plant parasitic nematodes as invasive species: characteristics, uncertainty and biosecurity implications. Annals of Applied Biology 163:323–50.
  32. Starr, J. L. , Koenning, S. R. , Kirkpatrick, T. L. , Robinson, A. F. , Roberts, P. A. and Nichols, R. L. 2007. The future of nematode management in cotton. Journal of Nematology 39:283–294.
  33. Taylor, A. L. and Sasser, J. N. 1978. Biology, identification and control of root-knot nematodes North Carolina State University, Raleigh.
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FIGURES & TABLES

Figure 1:

Average reproduction factor of Meloidogyne species, multiplied in I. grandifolia, I. hederifolia, I. nil, I. purpurea, and I. quamoclit. *Means followed by the same letter in the columns do not differ by the Scott-Knott test at 5% probability.

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REFERENCES

  1. Anwar, S. S., Zia, A. and Javed, N. 2009. Meloidogyne incognita infection of five weed genotypes. Pakistan Journal of Zoology 41:95–100.
  2. Azania, C. A. M. , Marques, R. P. , Azania, A. A. P. M. and Rolim, J. C. 2009. Overcoming dormancy of Ipomoea quamoclit and I. hederifolia seeds. Planta Daninha 27:23–27.
  3. 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.
  4. Bellé, C. , Kulczynski, S. M. , Kaspary, T. E. and Kuhn, P. R. 2017b. Weeds as host alternative to Meloidogyne incognita. Nematropica 47:26–33.
  5. Bellé, C. , Kaspary, T. E. , Balardin, R. R. , Ramos, R. F. and Antoniolli, Z. I. 2019. Meloidogyne species associated with weeds in Rio Grande do Sul. Planta Daninha 37:e019214250.
  6. Bellé, C. , Ramos, R. F. , Balardin, R. R. , Nora, D. D. and Kaspary, T. E. 2020. Multiplication potential of Meloidogyne arenaria in weeds found in Brazil. European Journal of Plant Pathology 157:441–447.
  7. 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.
  8. Carneiro, R. M. D. G. , Monteiro, J. M. S. , Silva, U. C. and Gomes, G. 2016. “Gênero Meloidogyne: diagnose através de eletroforese de isoenzimas e marcadores SCAR”, In Oliveira, C. M. , Santos, M. A. and Castro, L. H. S. (Eds), Diagnose de fitonematoides Millennium, Campinas, pp. 71–93.
  9. Carvalho, S. J. P., López-Ovejero, R. F. and Christoffoleti, P. J. 2008. Growth and development of five weed species of the Amaranthus genus. Bragantia 67:317–.
  10. Chauhan, B. S. and Abugho, S. B. 2012. Threelobe morningglory (Ipomoea triloba) germination and response to herbicides. Weed Science 60:199–204.
  11. Cruz, C. D. 2006. Programa GENES: biometria UFV, Vicosa.
  12. Elmore, C. D. , Hurst, H. R. and Austin, D. F. 1990. Biology and control of morning-glories (Ipomoea spp.). Weed Science 5:83–114.
  13. Ferraz, L. C. C. B. , Pitelli, R. A. and Furlan, V. 1978. Nematoides associados a plantas daninhas na região de Jaboticabal, SP-primeiro relato. Planta Daninha 1:5–11.
  14. Ferreira, P. P. A. and Miotto, S. T. S. 2009. Synopsis of Ipomoea L. (Convolvulaceae) species in Rio Grande do Sul State, Brazil. Revista Brasileira de Biociências 7:440–453.
  15. Galbieri, R. , Davis, R. F. , Scoz, L. B. , Belot, J. L. and Skantar, A. M. 2020. First Report of Meloidogyne enterolobii on Cotton in Brazil. Plant Disease 104:2295, available at: https://doi.org/10.1094/PDIS-02-20-0365-PDN.
  16. Groth, M. Z , Bellé, C. , Cocco, K. L. T. , Kaspary, T. E. , Casarotto, G. , Cutti, L. and Schmitt, J. 2017. First report of Meloidogyne enterolobii infecting the weed Jerusalem cherry (Solanum pseudocapsicum) in Brazil. Plant Disease 101:510.
  17. Heap, I. 2020. The International Survey of Herbicide Resistant Weeds, available at: http://www.weedscience.org.
  18. Holm, L. , Doll, J. , Holm, E. , Pancho, J. and Herberger, J. 1997. World weeds: natural histories and distribution Wiley, New York.
  19. 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.
  20. Kaspary, T. E. , Bellé, C. , Groth, M. Z. , Cocco, K. L. T. , Cutti, L. and Casarotto, G. 2017. Amaranthus viridis is a weed host of Meloidogyne arenaria in Rio Grande do Sul State, Brazil. Plant Disease 101:635.
  21. Kissmann, K. G. and Groth, D. 1999. “Convolvulaceae Juss, Vol. 1”, In Kissmann, K. G. and Groth, D. (Eds), Plantas infestantes e nocivas 2nd ed., BASF, São Paulo, pp. 673–693.
  22. Koenning, S. R. , Overstreet, C. , Noling, J. W. , Donald, P. A. , Becker, J. O. and Fortnum, B. A. 1999. Survey of crop losses in response to phytoparasitic nematodes in the United States for 1994. Journal of Nematology 31:587–618.
  23. Lorenzi, H. 2000. Plantas daninhas do Brasil: terrestres, aquáticas, parasitas e tóxicas Instituto Plantarum, Nova Odessa.
  24. McSorley, R. 1992. Nematological problems in tropical and subtropical fruit tree crops. Nematropica 22:103–116.
  25. Moens, M. and Perry, R. 2009. Migratory plant endoparasitic nematodes: a group rich in contrasts and divergence. Annual Review Phytopathology 37: 313–32.
  26. Oostenbrink, M. 1966. Major characteristics of the relation between nematodes and plants. Mendelingen Landbouwhogeschool, Wageningen 66:1–46.
  27. Ramesh, K. , Matloob, A. , Aslam, F. , Florentine, S. K. and Chauhan, B. S. 2017. Weeds in a changing climate: vulnerabilities, consequences, and implications for future weed management. Frontiers in Plant Science 8:95.
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