Differences in parasitism of root-knot nematodes (Meloidogyne spp.) on oilseed radish and oat

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Differences in parasitism of root-knot nematodes (Meloidogyne spp.) on oilseed radish and oat

Negin Hamidi / Abolfazl Hajihassani *

Keywords : Host response, histopathology, oilseed radish, oat, root-knot nematodes, resistance

Citation Information : Journal of Nematology. Volume 52, Pages 1-10, DOI: https://doi.org/10.21307/jofnem-2020-043

License : (CC-BY-4.0)

Received Date : 02-January-2020 / Published Online: 24-April-2020

ARTICLE

ABSTRACT

Oilseed radish and oat are cool season annual crops that are potentially used as “trap” or “biofumigant” crops for the suppression of plant-parasitic nematodes in soil. Cultivars of oilseed radish (Carwoodi, Cardinal, Final, Image, Concorde, Control, Eco-Till, Karakter and Cannavaro), white (Tachiibuki) and black (Pratex) oats were evaluated for their ability to reduce reproduction of three root-knot nematode species: Meloidogyne javanica, M. incognita race 3, and M. arenaria race 1. Nematode penetration and development were also evaluated using selected resistant and susceptible cultivars under greenhouse conditions. Root galling severity, number of eggs per gram of fresh root, and rate of reproduction varied among the cultivars in response to nematode infection. Oilseed radish cv. Carwoodi was resistant to M. javanica, whereas Karakter and Concorde were maintenance hosts allowing the nematode to maintain or increase its population on the plants. For M. incognita, Control and Carwoodi oilseed radish and Tachiibuki oat were resistant hosts. The cultivars that supported little reproduction of M. arenaria were Karakter and Carwoodi radish, and Tachiibuki oat. Comparable numbers of nematodes entered the roots of susceptible and resistant cultivars of oilseed radish and oat during early stages of infection. However, the development of the nematodes as evident from counting young and egg-laying females in roots were significantly decreased or inhibited in the resistant cultivars compared to the susceptible cultivars indicating that resistance occurs at post-infection stages. Histopathological examinations of galled-root tissues also revealed the susceptibility and resistance responses of selected cultivars of oilseed radish and oat to these nematode species.

Graphical ABSTRACT

Root-knot nematodes (Meloidogyne spp.) are obligate, sedentary endoparasites that can infect both the below- and above-ground parts of many crops at different developmental stages. Meloidogyne spp. are serious pathogens of multiple vegetable crops in the southern United States and worldwide (Johnson et al., 1992; Jones et al., 2013; Hajihassani, Davis, and Timper, 2019; Hajihassani, Rutter, and Luo, 2019). In Georgia, more than 66% of vegetable growing areas are infested with at least one species of root-knot nematodes at the level of one nematode per 100 cm3 of soil where management is needed to achieve economically reasonable yields (Marquez and Hajihassani, 2019). The nematode species most associated with vegetable production in Georgia are M. incognita, M. arenaria, and M. javanica (Marquez et al., 2019).

The life cycle of root-knot nematodes is completed in three basic stages: egg, juvenile (J), and adult stages. Eggs are deposited in a gelatinous mass that protects eggs from environmental extreme (e.g. high temperature) and microbes. The egg masses can be found on the surface of galled roots or embedded within the gall tissue. Under favorable conditions, egg development takes place and the first stage juvenile (J1) is formed. The J1 molts into J2 stage that hatch from eggs. At the onset of parasitism, the infective J2s penetrate the roots and migrate toward the vascular system of root tissues where they develop permanent feeding sites termed “giant cells.” This feeding site serves as a nutrient source for the growth of J2 and other development stages. The J2 molts to become J3, which develops into J4 stage and then adult females which lay eggs in a gelatinous sac (Caillaud et al., 2008; Jones et al., 2013).

For decades, pre-planting treatment of soil beds with fumigant chemicals has been the primary practice for controlling nematodes in vegetable crops in the southern United States; however, use of fumigants is being restricted because of their broad-spectrum and notoriously toxic effects (Zasada et al., 2010). The phase out of methyl bromide, and high costs and application difficulties associated with other chemical fumigants have led to identification and development of potential alternatives for management of Meloidogyne spp. (Hajihassani, Davis, and Timper, 2019; Hajihassani, Rutter, and Luo, 2019).

In recent decades, Brassica crops have been recognized for control of root-knot nematodes based on their potential use in biofumigation approaches through producing biocidal compounds such as isothiocyanates, and their ability to inhibit the development and reproduction of nematodes (Johnson et al., 1992; Kirkegaard and Sarwar, 1998; Edwards and Ploeg, 2014). Likewise, cereal cover crops including oats can act as non-hosts or poor hosts to Meloidogyne spp., resulting in reduced nematode numbers in soil (Wang et al., 2004), if used in a proper crop sequence program. Oilseed radish (Raphanus sativus var. oleiformis), white (Avena staiva) and black oat (Avena strigosa) are cool season annual crops that can be grown in the fall and harvested in the spring in regions with mild winter temperatures. These plant species are considered as potential “trap” or “biofumigant” crops for suppressing nematode populations in soil in an integrated program to lessen the use of fumigant nematicides. However, many of oilseed radish and oat cultivars are susceptible to Meloidogyne spp. resulting in an undesired population increase in soil during the growth of the crops. To avoid this, oilseed radish and oats cultivars that are resistant should be identified.

Screening for resistance to Meloidogyne spp. in radish (Teklu et al., 2014; O’Neill, 2016; Waisen, 2019) and oat (Tateishi et al., 2011; De Brida et al., 2017; Uesugi et al., 2018) has been investigated and several genotypes (lines) and/or commercial cultivars have been characterized. Research on these crops has led to variable suppression of nematodes, depending upon the plant cultivar and nematode species examined (Steddom et al., 2008; Tateishi et al., 2008; Waisen, 2019). Limited information is available on the response of commercially available oilseed radish and oat cultivars toward major species of M. javanica, M. incognita, and M. arenaria. The purpose of this study was to evaluate the host suitability of oilseed radishes and oats for these nematode species, and assess the extent of variation in nematode penetration and development using known susceptible and resistant cultivars.

Materials and methods

Nematode inoculum

Pure cultures of M. javanica, M. incognita race 3, or M. arenaria race 1 were increased on eggplant (cv. Black Beauty) grown in steamed field soil in the greenhouse for two to three months. The nematodes were identified to species using species-specific primers and mitochondrial genes (Marquez et al., 2019). Inoculum for experiments was obtained by recovering second-stage juveniles (J2) of each nematode species from infected roots of eggplant by incubating chopped roots in a misting chamber for five to seven days. The J2 collected from a 38-µm-aperture sieve were stored at 5°C until use. The inoculation densities of J2 for the following experiments were determined from three replicates of 1 ml aliquots of the nematode suspension.

Plant materials

Nine cultivars of oilseed radish, and one cultivar each of white and black oat were evaluated. Seed of oilseed radish cv. Carwoodi, Cardinal, Final, and Image as well as Tachiibuki oat were obtained from Grassland Oregon, Salem, OR. Also, oilseed radish cv. Concorde, Control, Eco-Till, Karakter and Cannavaro as well as Pratex black oat were obtained from Allied Seed, LLC, Nampa, ID. Tomato cv. Rutgers, an excellent host for all root-knot nematode species, was included in the experiments as a standard host.

Host suitability assay

This study was conducted from June to October 2018. Seeds of the oilseed radish and oat cultivars were germinated in Miracle-Gro Moisture Control potting mix (The Scotts Miracle-Gro Company, Marysville, OH) in seed trays (Speedling Incorporated, Ruskin, FL). Two-week-old seedlings were transplanted individually into Deepot D40L cells (6.9-cm-diam. × 25.4-cm deep; Stuewe & Sons, Inc., Tangent, Oregon) filled with equal parts (v/v) of steamed field soil and washed sand. After two days, initial inoculum of M. javanica, M. incognita, and M. arenaria were applied to plants by pipetting 1,000 J2 into two depressions made in the soil at the base of the stems. The deepot cells were arranged in a completely randomized design with five replications on support trays (Stuewe & Sons, Inc., Tangent, OR), and plants were maintained for eight week, the time needed for the completion of two generations of Meloidogyne spp. Temperature in the greenhouse varied between 25 and 28°C. Plants were watered daily and fertilized once, at two-weeks-after planting with 2-g Osmocote smart-release fertilizer (15-9-12, The Scotts, Marysville, OH). Two separate experiments were conducted for each Meloidogyne species.

At the termination of each experiment, shoots of all plants were removed and root systems were washed free of soil. Root systems of each plant were weighed and assessed for severity of galling on a scale 0 to 5 where 0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; and 5 = more than 100 galls (Taylor and Sasser, 1978). Root systems from each plant were cut into 2-cm pieces, and eggs were extracted with a 1.0% NaOCL solution (Hussey and Barker, 1973) followed by the standard centrifugal flotation technique. Eggs were counted using an inverted microscope and total number of eggs per gram of fresh root was determined. The reproductive factor (Rf) of Meloidogyne spp. was assessed by dividing the final nematode counts extracted from roots by initial number of J2 inoculated to plants at the beginning of experiment, and used as a measure for the host suitability of oilseed radish and oat cultivars to Meloidogyne spp. Host suitability was evaluated as follows: Rf < 0.1, nonhost (highly resistant); 0.1 ≤ Rf < 1, poor host (resistant); 1 ≤ Rf < 2, maintenance host (susceptible); and Rf ≥ 2, good host (highly susceptible).

Nematode penetration and development assay

This assay was conducted from March to May 2019 using Carwoodi and Eco-Till oilseed radish and Tachiibuki and Pratex oat based on their response to Meloidogyne spp. in the host suitability study (Table 1). The seeds were planted in 72-cell plug flats (Johnny’s Selected Seeds, Fairfield, Maine) filled with steamed Miracle-Gro potting mix two to three weeks prior to nematode inoculation. Each two-week-old seedling was inoculated with 200 J2 of M. incognita race 3, M. arenaria race 1, or M. javanica in 1 ml water pipetted into two holes (2-cm deep) made around the stem. The experiment was a completely randomized design with three replicates for each cultivar. It was conducted in a greenhouse at 25±3°C. The plants were watered lightly each day. The infected root systems were collected one, three, and five days after inoculation (DAI) to assess nematode penetration. Three seedlings were randomly selected, washed gently with tap water to remove the soil, soaked in 0.5% NaOCL for 30 sec and then rinse with tap water. The nematode J2 in intact roots were stained by boiling for 30 sec in 30 ml tap water containing 1 ml of 3.5% (wt/vol) acid fuchsin. After staining, the roots were rinsed with tap water and destained by boiling in acidified glycerol for about 1 min. Visualization of stained nematodes was done by placing the root systems in a Petri dish and examination under a stereomicroscope (Stemi 508, ZEISS, Hannover, Germany). The penetration of Meloidogyne spp. was examined by counting the total number of J2 stained inside the roots at each sampling day. To examine nematode development and reproduction, three root systems were collected at 28 DAI, stained as specified previously and 20 galls were randomly examined under a stereomicroscope to count females. Swollen females with no egg-mass in the root tissues were considered young females, whereas egg-laying females were considered adult females (Fig. 2). This experiment was conducted twice.

Table 1.

Reproduction of Meloidogyne javanica, M. incognita race 3, and M. arenaria race 1 on oilseed radish, white (Tachiibuki) and black (Pratex) oats under greenhouse conditions.

10.21307_jofnem-2020-043-t001.jpg

A histopatological examination was also conducted to evaluate the response of Carwoodi and Eco-Till radish and Tachiibuki and Pratex oat to nematode infection. Two to three segments 1 to 2 cm long of galled roots from each infected plant were examined at 28 DAI. Root segments, including individual small galls and adjoining portions of non-galled tissue, were fixed in buffered formaldehyde solution for 24 to 48 hr at room temperature. The fixed segments were dehydrated in a graded ethanol series (70 to 100%) using a tissue processor (Excelsior AS, Thermo Scientific, Thermo Shandon Limited, Runcorn Cheshire, UK), and embedded in paraffin. Cross sections 4 to 5 µm thick were prepared using a rotary microtome (RM2125 RTS, Leica Biosystems, Nussloch, Germany), and then mounted on glass slides. The sections were stained with Hematoxylin-Eosin (HE), evaluated microscopically at 40 to 100 × magnifications (Olympus BX43, Olympus, Tokyo, Japan) and images captured using a digital camera (Olympus DP73, Olympus, Tokyo, Japan) (Hajihassani et al., 2020).

Statistical analysis

Galling severity, egg counts, and Rf from the host suitability test were subjected to one-way ANOVA, while data from the penetration and development assay were analyzed by two-way ANOVA using the PROC mixed models (SAS v. 9.4, SAS Institute, Cary, NC). Data were combined prior to ANOVA because the initial and repeat trials were not significantly different (P > 0.1). Plant cultivar and DAI (penetration and development assay) were considered fixed effects, and trial was a random effect. Means were compared using Tukey’s adjustment for multiple comparison test at P < 0.05. Least-squares means (LS-means) t-tests demonstrating if the mean was significantly different from 0 were used to determine whether the nematode Rf was different from 1 (the base 10 algorithm of 1 is 0). Therefore, single t-tests that showed the treatment mean was significantly different from 0 indicated that the Rf deviated significantly from 1 (Hajihassani et al., 2020).

Results

Host suitability of oilseed radish and oat for Meloidogyne species

Cultivar affected (P < 0.0001) all resistance characteristics including gall severity and nematode reproduction. Meloidogyne javanica, M. incognita, and M. arenaria produced varying numbers of galls and eggs in the roots of oilseed radish and oat cultivars (Fig. 1); however, significant differences among the cultivars were infrequent. Root galling severity on the oilseed radish and oat cultivars was in the range of 1.4 (Carwoodi oilseed radish) to 3.7 (Cannavaro radish) for M. javanica, 0.4 (Carwoodi) to 2.7 (Image radish and Pratex oat) for M. incognita, and 0.3 (Tachiibuki oat) to 3.3 (Eco-Till and Final) for M. arenaria. The numbers of M. javanica eggs per gram of fresh root ranged from 270 for Carwoodi to 4,946 for Cannavaro. Egg production by M. incognita varied from 21 for Carwoodi to 2,436 eggs per gram root for Image. For M. arenaria, Tachiibuki (32) and Eco-Till (3,780) had the minimum and maximum number of eggs per gram of root (Fig. 1). All three nematode species reproduced more on tomato than on the oat cultivars and most of the oilseed radish cultivars (Table 1). Meloidogyne javanica reproduced on eight cultivars of oilseed radish, and the white and black oats. Although the nematode had an Rf of 1.05 on Carwoodi, the Rf was not different from 1 based on the LS-means t-tests indicating that M. javanica failed to increase its population on Carwoodi.

Figure 1:

Gall and egg formation of Meloidogyne javanica (A and B), M. incognita race 3 (C and D), and M. arenaria race 1 (E and F) on cultivars of oilseed radish, white (Tachiibuki) and black (Pratex) oats under greenhouse conditions. Tomato cv. Rutgers was used as a susceptible host. Gall severity was assessed based on a scale 0 to 5 where 0 = no galls, and 5 ≥ 100 galls on roots. Data are the mean of two trials with ten replicates. Bars with the same letter(s) are not significantly different based on the Tukey’s test.

10.21307_jofnem-2020-043-f001.jpg

The Rf of the three nematode species did not often vary among the cultivars tested. The Rf of M. incognita was lower (P < 0.05) in Carwoodi and Control oilseed radish and Tachiibuki oat than all other cultivars examined. Meloidogyne javanica and M. incognita exhibited only slight reproduction on Karakter and Concorde, having Rf’s ranging from 1.0 to 1.9. The Rf of M. arenaria was significantly less than 1 on Karakter and Carwoodi oilseed radish and Tachiibuki oat compared to all other cultivars.

Penetration and development of Meloidogyne spp.

The penetration of J2 into roots of oilseed radish and oat was only affected (P < 0.0001) by DAI, showing that the susceptibility or resistance of the cultivars had no effect on nematode root penetration. At 1 DAI, J2 were observed in the roots of oilseed radish; however, no difference in the numbers of J2 was observed between susceptible Eco-Till and resistant Carwoodi cultivars for M. javanica, M. incognita, or M. arenaria (Table 2). A similar trend in the penetration of the nematodes in roots of susceptible Pratex and resistant Tachiibuki oats was also detected (Table 3). At 3 and 5 DAI, comparable numbers of J2 penetrated the roots of susceptible and resistant cultivars of oilseed radish and oat. Nematode development was studied by counting the numbers of young and adult (egg-laying) females of each nematode species within galled (swelled) roots at 28 DAI. For all three nematode species, greater numbers (P < 0.05) of young and adult females were observed in the root tissues of the susceptible cultivar Eco-Till than in resistant Carwoodi (Table 2). Similarly, the total number of young and adult females for M. incognita and M. arenaria was higher (P < 0.05) in the roots of susceptible Pratex (Fig. 2A and B) compared to resistant Tachiibuki oat (Table 3).

Table 2.

Number of second-stage juveniles (J2), and young and adult (egg-laying) females of Meloidogyne spp. in roots of oilseed radish (Eco-Till, susceptible; Carwoodi, resistance) at early-stage of infection (1, 3, and 5 days after inoculation; DAI) and at 28 DAI in the greenhouse.

10.21307_jofnem-2020-043-t002.jpg
Table 3.

Number of second-stage juveniles (J2), and young and adult (egg-laying) females of Meloidogyne spp. in roots of oat (Pratex, susceptible; Tachiibuki, resistance) cultivars at early-stage of infection (1, 3, and 5 days after inoculation; DAI) and at 28 DAI in the greenhouse.

10.21307_jofnem-2020-043-t003.jpg
Figure 2:

Representative images of roots of oilseed radish cv. Eco-Till (A) and oat cv. Pratex (B) infected with Meloidogyne arenaria at 28 days after inoculation.

10.21307_jofnem-2020-043-f002.jpg

Cross sections of the root system of oilseed radish and oat infected with all three root-knot nematode species showed morphological variations between susceptible and resistant cultivars. At 28 DAI, more than 96% of the root tissues of resistant Carwoodi (Fig. 3A) and Tachiibuki (not shown) inoculated with M. incognita or M. arenaria had no sign of feeding site or giant cell formation. In contrast, all the root tissues of susceptible Eco-Till (Fig. 3B) and Pratex (not shown) infected with M. arenaria or M. incognita had multiple giant cells. We noticed very limited hypertrophy in cortical cells and the vascular cylinder of Carwoodi and Tachiibuki infected with these two nematode species, suggesting that the nematode J2 penetrated the roots but failed to establish giant cells. Multinucleated giant cells were detected in 100% of the root tissues of Eco-Till infected with M. javanica (not shown). Cross sections of the root system of M. javanica-infected Carwoodi showed degraded giant cells in 5 to 10% of the roots (not shown). Multiple giant cells and severe hypertrophy in the vascular cylinder and cortical cells were detected in the root tissues of Tachiibuki and Pratex oats infected with M. javanica (Fig. 4A, B).

Figure 3:

Hematoxylin-Eosin stained cross sections of root tissues of oilseed radish cv. Carwoodi (A) and Eco-Till (B) infected with Meloidogyne incognita at 28 days after inoculation. N, female nematode; GC, giant cell.

10.21307_jofnem-2020-043-f003.jpg
Figure 4:

Cross sections of root tissues of oat cv. Pratex (A) and Tachiibuki (B) infected with Meloidogyne javanica at 28 days after inoculation. N, female nematode; GC, giant cell.

10.21307_jofnem-2020-043-f004.jpg

Discussion

Management of plant-parasitic nematodes in vegetable plasticulture or bare-ground systems in the southern US relies heavily on pre-plant fumigation of soil with 1,3-dichloropropene (Telone II), Chloropicrin or metam sodium (Vapam), followed by in-season application of non-fumigant nematicides (Hajihassani, Davis, and Timper, 2019). One of the alternative management methods is the use of nematode-suppressive cover/trap crops to reduce nematode populations in soil. However, the majority of oilseed and oat cultivars have variable levels of susceptibility to root-knot nematodes, and characterization of resistant cultivars could have important implications in root-knot nematode control. Results from this study suggest that differential responses to infection with M. javanica, M. incognita, and M. arenaria exist in oilseed radish and oat cultivars. We determined the susceptibility/resistance of representative oilseed radish and oat cultivars for each nematode species by comparing the nematode Rf. Other resistance characteristics such as root galling severity and low recovery of eggs from root tissues also indicate that only a few cultivars of oilseed radish and oats evaluated exhibited resistance relative to the susceptible tomato.

In this study, M. javanica greatly reproduced on most of the oilseed radish and oat cultivars, and only Carwoodi radish was resistant to M. javanica. The level of resistance to M. incognita varied among oilseed radish and oat cultivars. Control and Carwoodi radish and Tachiibuki oat did not support reproduction and were resistant to M. incognita. Results from this study for oilseed radish are consistent with previous studies (Waisen, 2019) reporting several cultivars of oilseed radish are poor hosts to M. incognita. We found that the Rf of M. javanica and M. incognita on Karakter and Concorde radish was slightly greater than 1 and these two cultivars can retain or increase the nematode populations in soil. M. arenaria did not infect Karakter and Carwoodi radish and Tachiibuki oat. All other cultivars were susceptible or highly susceptible to the nematode. A greenhouse study in Washington has shown that Carwoodi oilseed radish is resistant to M. chitwoodi (O’Neill, 2016). Our results confirmed that this cultivar is also resistant to M. javanica, M. incognita and M. arenaria. Our data on Tachiibuki white oat are consistent with previous research in which few or no eggs of M. incognita and M. arenaria were detected in the roots of Tachiibuki (Tateishi et al., 2011). Uesugi et al. (2018) reported that a black oat line KH1a significantly reduced the reproduction of different isolates of M. incognita, M. arenaria, M. javanica, and M. hapla. In the present study, all three species of M. incognita, M. arenaria, and M. javanica were pathogenic on Pratex black oat.

Resistance to root-knot nematodes can occur prior to infection by reducing nematode invasion and/or penetration into host plants, at post-infection by limiting or containing nematode movement, development, and reproduction in root tissues, and at both the pre- and post-infection stages (Trudgill, 1991; Proite et al., 2008; Hajihassani, Rutter and Luo, 2019). To investigate the initial infection process of three species of root-knot nematodes in oilseed radish and oat, the penetration of J2 into root systems of selected susceptible and resistant cultivars were examined at 1, 3, and 5 DAI. We found no significant differences in the penetration of M. javanica, M. incognita, and M. arenaria between susceptible and resistant cultivars, suggesting that the resistance to these nematodes does not occur at early stages of infection. Tateishi et al. (2011) reported that similar numbers of M. incognita, M. arenaria, and M. hapla penetrated the resistant Tachiibuki oat and a susceptible host. In contrast, greater numbers of young and mature egg-laying females for all nematode species were observed in infected roots of susceptible Eco-Till radish and Pretext oat at 28 DAI, demonstrating that the resistance could act at post-infection stages. In previous research, roots of Tachiibuki oat infected with M. incognita or M. arenaria had a lower number of mature females compared to a susceptible host at 30 DAI (Tateishi et al., 2011).

The histopathological observations in our study showed that the root structure in the Meloidogyne-infected resistant cultivars of oilseed radish and oat were healthy with no or limited signs of feeding site development. The absence of nematodes and giant cells in the resistant cultivars may be attributed to inhibition of juvenile development in the root tissues. In contrast, the root tissues of susceptible oilseed and oat cultivars had several hypertrophied and multinucleated cells due to root-knot nematode development (Cabasan et al., 2014; Hajihassani et al., 2020). Giant cells serve as an important source of nutrients for nematode growth and reproduction. These structures restrict the translocation of water and nutrients from nematode-infected roots to aboveground parts of the plant (Hussey and Williamson, 1997; Caillaud et al., 2008), thereby adversely affecting plant growth.

In summary, the results suggest that development and reproduction of M. javanica are reduced on Carwoodi oilseed radish; M. incognita on Control and Carwoodi radishes, and Tachiibuki oat; and M. arenaria on Carwoodi and Karakter radishes, and Tachiibuki oat. Crop rotations with these resistant cultivars could be beneficial in control of these yield-limiting nematode species. Oilseed radish and oat are not only grown as cash crops but their potential use as suppressive “biofumigant” and/or “trap” crops for nematode management is of great importance. However, special attention to presence of other plant-parasitic nematodes in soil is necessary when cover cropping with oilseed radishes or oats. For example, Grabau et al. (2017) reported that Defender oilseed radish can increase populations of the root-lesion nematode Pratylenchus penetrans in soil (Grabau et al., 2017). Pratex oat which was susceptible to all three root-knot nematode species in this study can have positive impact on reducing stubby-root-nematode (Paratrichodorus spp.) populations (USDA-NRCS, 2014). The efficacy of these winter-grown cover crops alone or in combination with in-season use of non-fumigant nematicides for the control of plant-parasitic nematodes in vegetable systems are under investigation in field trials in Georgia, USA.

Acknowledgements

The authors greatly appreciate Grassland Oregon, Salem, OR and Allied Seed, LLC., Nampa, ID for supplying the seed of oilseed radishes and oats.

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  12. Jones, J. T. , Haegeman, A. , Danchin, E. G. , Gaur, H. S. , Helder, J. , Jones, M. G. , Kikuchi, T. , Manzanilla-Lopez, R. , Palomares-Rius, J. E. , Wesemael, W. M. and Perry, R. N. 2013. Top 10 plant-parasitic nematodes in molecular plant pathology. Molecular Plant Pathology 14:946–961.
  13. Kirkegaard, J. A. and Sarwar, M. 1998. Biofumigation potential of Brassicas 1. Variation in glucosinolate profiles of diverse field grown brassicas. Plant and Soil 201:71–89.
  14. Marquez, J. and Hajihassani, A. 2019. Incidence, abundance and distribution of plant-parasitic nematodes in vegetable systems of southern Georgia. Journal of Nematology 51:28 (Abstr.).
  15. Marquez, J. , Ye, W. , Skantar, A. and Hajihassani, A. 2019. Molecular identification of Meloidogyne spp. associated with vegetable crops in southern Georgia. Journal of Nematology 51:29 (Abstr.).
  16. O’Neill, K. P. J. 2016. “Cover cropping for control of Columbia root knot nematodes in short season potato production”, MS dissertation, Washington State University, Pullman, WA.
  17. Proite, K. , Carneiro, R. , Falcão, R. , Gomes, A. , Leal-Bertioli, S. , Guimarães, P. and Bertioli, D. 2008. Post-infection development and histopathology of Meloidogyne arenaria race 1 on Arachis spp. Plant Pathology 57:974–980.
  18. Steddom, K. , Ong, K. and Starr, J. 2008. Efficacy of various brassica varieties for the suppression of root knot, ring, and stunt nematodes. Phytopathology 98:S150.
  19. Tateishi, Y. , Sano, Z. , Iwahori, H. , Uesugi, K. , Katsura, M. and Gau, M. 2008. Suppression of root-knot nematode damage to succeeding sweet potato by summer-sown cultivation of an oat variety, ‘Tachiibuki’. Japanese Journal of Nematology 38:1–7.
  20. Tateishi, Y. , Iwahori, H. , Uesugi, K. and Katsura, M. 2011. Invasion, development, and reproduction of 3 Meloidogyne species on oat cultivar Tachiibuki, a nematode-suppressive fall crop. Nematological Research 41:1–7.
  21. Taylor, A. L. and Sasser, J. N. 1978. Biology, identification, and control of root-knot nematodes (Meloidogyne species). North Carolina State University Graphics, Raleigh, NC.
  22. Teklu, M. G. , Schomaker, C. H. and Been, T. H. 2014. Relative susceptibilities of five fodder radish varieties (Raphanus sativus var. oleiformis) to Meloidogyne chitwoodi . Nematology 16:577–590.
  23. Trudgill, D. L. 1991. Resistance to and tolerance of plant parasitic nematodes in plants. Annual Review of Phytopathology 29:167–193.
  24. USDA-NRCS 2014. Black oat (Avena strigose Schreb), available at: https://plants.usda.gov/plantguide/pdf/pg_avst2.pdf (Accessed March 26, 2020).
  25. Uesugi, K. , Katsura, M. , Uwatoko, N. , Tateishi, Y. , Murata, G. and Iwabuchi, K. 2018. Suppressive effect of black oat, Avena strigosa, KH1a on Meloidogyne spp. Nematology 20:387–396.
  26. Wang, K. H. , McSorley, R. and Gallaher, R. N. 2004. Effect of winter cover crops on nematode population levels in north Florida. Journal of Nematology 36:517–523.
  27. Waisen, P. 2019. Management of parasitic nematodes and soil health using oil radish (Raphanus sativus) and brown mustard (Brassica juncea) cover crops. PhD dissertation, University of Hawaii, Manoa, HI.
  28. Zasada, I. A. , Halbrendt, J. M. , Kokalis-Burelle, N. , LaMondia, J. , McKenry, M. V. and Noling, J. W. 2010. Managing nematodes without methyl bromide. Annual Review of Phytopathology 48:311–328.
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FIGURES & TABLES

Figure 1:

Gall and egg formation of Meloidogyne javanica (A and B), M. incognita race 3 (C and D), and M. arenaria race 1 (E and F) on cultivars of oilseed radish, white (Tachiibuki) and black (Pratex) oats under greenhouse conditions. Tomato cv. Rutgers was used as a susceptible host. Gall severity was assessed based on a scale 0 to 5 where 0 = no galls, and 5 ≥ 100 galls on roots. Data are the mean of two trials with ten replicates. Bars with the same letter(s) are not significantly different based on the Tukey’s test.

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Figure 2:

Representative images of roots of oilseed radish cv. Eco-Till (A) and oat cv. Pratex (B) infected with Meloidogyne arenaria at 28 days after inoculation.

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Figure 3:

Hematoxylin-Eosin stained cross sections of root tissues of oilseed radish cv. Carwoodi (A) and Eco-Till (B) infected with Meloidogyne incognita at 28 days after inoculation. N, female nematode; GC, giant cell.

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Figure 4:

Cross sections of root tissues of oat cv. Pratex (A) and Tachiibuki (B) infected with Meloidogyne javanica at 28 days after inoculation. N, female nematode; GC, giant cell.

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REFERENCES

  1. Cabasan, M. T. N. , Umar, A. , Bellafiore, S. and De Waele, D. 2014. Histopathology of the rice root-knot nematode, Meloidogyne graminicola, on Oryza sativa and O. glaberrima . Nematology 16:73–81.
  2. Caillaud, M. C. , Abad, P. and Favery, B. 2008. Cytoskeleton reorganization, a key process in root-knot nematode-induced giant cell ontogenesis. Plant Signaling & Behavior 3:816–818.
  3. De Brida, A. L. , Correia, E. C. , Castro, B. C. , Zanuncio, J. and Wilcken, S. R. 2017. Oat, wheat, and sorghum genotype reactions to Meloidogyne incognita and Meloidogyne javanica . Journal of Nematology 49:386–389.
  4. Edwards, S. and Ploeg, A. 2014. Evaluation of 31 potential biofumigant Brassicaceous plants as hosts for three Meloidogyne species. Journal of Nematology 46:287–295.
  5. Grabau, Z. J. , Maung, Z. T. Z. , Noyes, D. C. , Bass, D. G. , Werling, B. P., Brainard, D. C. and Melakeberhan, H. 2017. Effects of cover crops on Pratylenchus penetrans and the nematode community in carrot production. Journal of Nematology 49:114–123.
  6. Hajihassani, A. , Davis, R. F. and Timper, P. 2019. Evaluation of selected nonfumigant nematicides on increasing inoculation density of Meloidogyne incognita on cucumber. Plant Disease 103:3161–3165.
  7. Hajihassani, A. , Rutter, W. B. and Luo, X. 2019. Resistant pepper carrying N, Me1, and Me3 have different effects on penetration and reproduction of four major Meloidogyne species. Journal of Nematology 51:1–9.
  8. Hajihassani, A. , Rutter, W. B. , Shartz, T. , Woldemeskel, M. , Ali, M. E. and Hamidi, N. 2020. Characterization of resistance to major tropical root-knot nematodes (Meloidogyne spp.) in Solanum sisymbriifolium . Phytopathology, pp. 666–673.
  9. Hussey, R. S. and Barker, K. R. 1973. Comparison of methods of collecting inocula of Meloidogyne spp., including a new technique. Plant Disease Reporter 57:1025–1028.
  10. Hussey, R. S. and Williamson, V. M. 1997. “Physiological and molecular aspects of nematode parasitism”, in Barker, K. R. , Pederson., G. A. and Windham, G. L. (Eds), Plant and Nematode Interactions American Society of Agronomy, Madison, WI, pp. 87–108.
  11. Johnson, A. W. , Golden, A. M. , Auld, D. L. and Summer, D. R. 1992. Effects of rapeseed and vetch as green manure crops and fallow on nematodes and soilborne pathogens. Journal of Nematology 24:117–127.
  12. Jones, J. T. , Haegeman, A. , Danchin, E. G. , Gaur, H. S. , Helder, J. , Jones, M. G. , Kikuchi, T. , Manzanilla-Lopez, R. , Palomares-Rius, J. E. , Wesemael, W. M. and Perry, R. N. 2013. Top 10 plant-parasitic nematodes in molecular plant pathology. Molecular Plant Pathology 14:946–961.
  13. Kirkegaard, J. A. and Sarwar, M. 1998. Biofumigation potential of Brassicas 1. Variation in glucosinolate profiles of diverse field grown brassicas. Plant and Soil 201:71–89.
  14. Marquez, J. and Hajihassani, A. 2019. Incidence, abundance and distribution of plant-parasitic nematodes in vegetable systems of southern Georgia. Journal of Nematology 51:28 (Abstr.).
  15. Marquez, J. , Ye, W. , Skantar, A. and Hajihassani, A. 2019. Molecular identification of Meloidogyne spp. associated with vegetable crops in southern Georgia. Journal of Nematology 51:29 (Abstr.).
  16. O’Neill, K. P. J. 2016. “Cover cropping for control of Columbia root knot nematodes in short season potato production”, MS dissertation, Washington State University, Pullman, WA.
  17. Proite, K. , Carneiro, R. , Falcão, R. , Gomes, A. , Leal-Bertioli, S. , Guimarães, P. and Bertioli, D. 2008. Post-infection development and histopathology of Meloidogyne arenaria race 1 on Arachis spp. Plant Pathology 57:974–980.
  18. Steddom, K. , Ong, K. and Starr, J. 2008. Efficacy of various brassica varieties for the suppression of root knot, ring, and stunt nematodes. Phytopathology 98:S150.
  19. Tateishi, Y. , Sano, Z. , Iwahori, H. , Uesugi, K. , Katsura, M. and Gau, M. 2008. Suppression of root-knot nematode damage to succeeding sweet potato by summer-sown cultivation of an oat variety, ‘Tachiibuki’. Japanese Journal of Nematology 38:1–7.
  20. Tateishi, Y. , Iwahori, H. , Uesugi, K. and Katsura, M. 2011. Invasion, development, and reproduction of 3 Meloidogyne species on oat cultivar Tachiibuki, a nematode-suppressive fall crop. Nematological Research 41:1–7.
  21. Taylor, A. L. and Sasser, J. N. 1978. Biology, identification, and control of root-knot nematodes (Meloidogyne species). North Carolina State University Graphics, Raleigh, NC.
  22. Teklu, M. G. , Schomaker, C. H. and Been, T. H. 2014. Relative susceptibilities of five fodder radish varieties (Raphanus sativus var. oleiformis) to Meloidogyne chitwoodi . Nematology 16:577–590.
  23. Trudgill, D. L. 1991. Resistance to and tolerance of plant parasitic nematodes in plants. Annual Review of Phytopathology 29:167–193.
  24. USDA-NRCS 2014. Black oat (Avena strigose Schreb), available at: https://plants.usda.gov/plantguide/pdf/pg_avst2.pdf (Accessed March 26, 2020).
  25. Uesugi, K. , Katsura, M. , Uwatoko, N. , Tateishi, Y. , Murata, G. and Iwabuchi, K. 2018. Suppressive effect of black oat, Avena strigosa, KH1a on Meloidogyne spp. Nematology 20:387–396.
  26. Wang, K. H. , McSorley, R. and Gallaher, R. N. 2004. Effect of winter cover crops on nematode population levels in north Florida. Journal of Nematology 36:517–523.
  27. Waisen, P. 2019. Management of parasitic nematodes and soil health using oil radish (Raphanus sativus) and brown mustard (Brassica juncea) cover crops. PhD dissertation, University of Hawaii, Manoa, HI.
  28. Zasada, I. A. , Halbrendt, J. M. , Kokalis-Burelle, N. , LaMondia, J. , McKenry, M. V. and Noling, J. W. 2010. Managing nematodes without methyl bromide. Annual Review of Phytopathology 48:311–328.

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