Effects of vermicompost water extract prepared from bamboo and kudzu against Meloidogyne incognita and Rotylenchulus reniformis

Weed vermicompost made from moso-bamboo (Phyllostachys edulis (Carrière) J. Houz.) and Kudzu (Pueraria lobata (Willd) Ohwi) and vegetable vermicompost made from vegetable food waste were prepared as described by You et al. (2018) and Mishra et al. (2017), respectively. Weed vermicompost was prepared from 10 kg of the bamboo shoots powder mixed with 100 g of air dried kudzu vine pieces (< 10 cm in length) and 20 g of a commercial horse manure/wheat straw compost (Iris Ohyama Inc., Sendai, Japan) and adding 100 g of red wiggler (Eisenia fetida Savigny) (Commercial name ‘Kumataro-futomushi,’ Yokomizo-shokai Inc., Mito, Japan) in a closed container to conduct vermicomposting for 2  mon. Vegetable vermicompost was initiated 3 yr prior to this experiment with approximately 100 g of commercial mix of red wiggler (E. fetida) and blue worms (Perionyx excavatus Perrier) (Waikiki Worms Co., Honolulu, HI). The worms were fed weekly with vegetable food waste such as lettuce, kale, papaya, and banana peel. All earthworms were removed from the vermicompost right before VCT preparation. The VCTs were prepared fresh for each experiments, and were prepared by mixing each type of vermicompost in water at 1:10 (v/v) ratio, and aerated for 24 hr using 2.5 W Elite 800 air pumps (Rolf C. Hagen Inc., Montreal, Canada) at room temperature (24). The VCT was filtered using a kitchen strainer to separate the solid from the liquid prior to application. Samples of weed VCT and vegetable VCT were submitted to the Agriculture Diagnostic Services Center (ADSC) at the University of Hawaii at Manoa to assay for concentrations of macro-nutrients (nitrogen, phosphorus, potassium, calcium, magnesium, and boron) and micro-nutrients (Fe, Mn, Zn, and Cu).

A laboratory assay was conducted to examine the effects of weed VCT and vegetable VCT on the hatching of M. incognita and R. reniformis compared to that of water control. Meloidogyne incognita eggs were extracted from coleus (Plectranthus scutellarioides (L.) R. Br.) roots and R. reniformis eggs were extracted from pineapple (Ananas comosus (L.) Merr.) roots using NaOCl and centrifugal flotation methods (Hussey and Barker 1973). Water, weed VCT or vegetable VCT were contained in 60-ml plastic cups at 15 ml/cup. The experiment was arranged in complete randomized design with four replications. Each plastic cup served as a hatching chamber where approximately 200 freshly extracted M. incognita eggs or R. reniformis eggs were suspended in 200 μl of water over a 60.33-μm pore size screen (Fig. 1). This mesh size kept nematode eggs on the screen but allowed second stage juveniles (J2s) to pass through. To avoid bacterial growth, VCT or water was replaced every day with fresh VCT or water. Hatched J2s were collected every day for 7 d and total hatching was counted under an inverted microscope (Leica DMIL LED, Wetzlar, Germany). The egg hatching experiment was repeated once.

Figure 1

A laboratory assay was established to examine the effects of weed VCT and vegetable VCT on the mobility of M. incognita J2s. After adding 10 ml of water, weed VCT or vegetable VCT into individual 60-mm diameter petri plates, 0.5 ml of water suspension containing approximately 100 freshly hatched M. incognita J2s were added to each plate. Treatments were replicated four times and the experiments were repeated once. Mobility of M. incognita J2s was examined by probing with a dental probe after 24 hr incubation of the nematodes in the solutions. Percentile of immobilized nematodes was calculated for each petri dish.

Two greenhouse trials were conducted in the Gilmore Greenhouse at the University of Hawaii at Manoa, Honolulu, HI from April to July 2017 to examine the effects of the VCTs on root penetration of M. incognita on ‘Bush Champion’ cucumber (Cucumis sativus L.), and that of R. reniformis on ‘Blackeye #5’ cowpea (Vigna unguiculata (L.) Walp.) by inducing host-plant resistance against nematodes. Roots of cucumber or cowpea seedlings were split into two parts and transplanted into two conjoined pots (Fig. 2). The purpose of using split-root assay was to avoid direct contact of VCTs on the tested nematodes that would lead to parasitism or immobilization of the nematodes by the chemical compounds or microbes in the VCTs. One side of the root system was drenched with weed VCT, vegetable VCT, or water 3 d prior to inoculation of the targeted nematode (Fig. 2). Two hundred J2s of M. incognita or 100 J2s of R. reniformis were introduced into the untreated conjoined pot. One week after M. incognita inoculation, or 3 wk after R. reniformis inoculation, roots from the nematode inoculated side were stained with Acid Fuchsin (Daykin and Hussey, 1985) and quantified for nematode penetration under the microscope.

Figure 2

To examine biological activities of weed vermicompost or vegetable vermicompost, free-living nematodes were used as bioindicators. Nematodes were extracted from 30 cm³ weed vermicompost or vegetable vermicompost by immersing the vermicompost into 300 ml water using Baermann trays for 24 hr (Southey, 1986). Each treatment was replicated three times. Bacterivorous and omnivorous nematodes, the two most dominant nematode trophic groups present, were counted under an inverted microscope (Leica DMIL LED, Wetzlar, Germany).

Two field trials were conducted at the Poamoho Research Station in Waialua, Oahu, HI to compare the mitigation of nematode damage on ‘Black Eye #5’ cowpea by weed VCT or vegetable VCT compared to water control using cowpea as a bioassay crop in field naturally infested with M. incognita and R. reniformis. Cowpea plants were drenched with weed VCT or vegetable VCT, or water weekly at 50 ml per plant during the first 2 wk, followed by 250 ml per plant for the rest of the crop over a three-month growing period from 27 April to 12 July, 2017. Each experimental plot had eight cowpea plants in a 1 × 3 m² -area. The three treatments were arranged in randomized complete block design with four replications. Soil nematode population densities were monitored at pre-plant, and at 1 and 2 mon after planting. Shoot and root weights, and root-gall index (RGI) were measured from three plants randomly selected in each plot at 2 mon after planting. Root galling was rated using a root-gall index based on a scale of 0 to 5, where 0 = 0, 1 = 1–2, 2 = 3–10, 3 = 11–30, 4 = 31–100, and 5 ≥ 100 galls (Taylor and Sasser, 1978). Cowpea pods from five plants per plot were harvested and weighted weekly from 21 June to 12 July, 2017.

Soil samples were collected 1 and 2 mon after cowpea planting in both trials. Four 20-cm deep soil samples were collected from each plot and combined into one bag. Nematodes were extracted from 250-cm³ soil by elutriation and centrifugal floatation (Jenkins, 1964; Byrd et al., 1976). All nematodes extracted were identified and assigned to a trophic group of bacterivores, fungivores, omnivores, or predators (Yeates et al., 1993), but herbivores were identified to the genus level with the aid of the inverted microscope described above.

Differences in macro- and micro-nutrient content between the weed VCT and vegetable VCT were analyzed by Student’s t-test. The other data were checked for normality, nematode data were log transformed [log₁₀(x+1)] if needed and subjected to one-way analysis of variance (ANOVA) using SAS (SAS Inc., Cary, NC). Repeated measures of nematode abundance from the cowpea field experiment were subjected to homogeneity of variance test over time. If there was no significant interaction between sampling date and treatment effect, data were subjected to repeated measures analysis. Means were separated using Waller–Duncan k-ration (k = 100) t-test. Only true means were presented.

Weed VCT contained lower concentrations of nitrogen, phosphorus, potassium, boron, Fe, and Cu than vegetable VCT, but both VCTs contained similar concentration of calcium, magnesium, Mn, and Zn levels (Table 1).

Both weed VCT and vegetable VCT suppressed M. incognita egg hatching compared to the water control (P ≤ 0.05, Fig. 3A). In the first trial of R. reniformis egg hatching test, both weed VCT and vegetable VCT reduced R. reniformis hatching compared to the water control. However, in the second trial, only weed VCT (P ≤ 0.05) reduced the hatching of R. reniformis compared to the water control (Fig. 3B).

Figure 3

Although vegetable VCT immobilized M. incognita J2s more effectively than weed VCT in Trial I (P ≤ 0.05), both VCTs suppress the J2s mobility equally in Trial II compared to the water control (P ≤ 0.05, Fig. 4).

Figure 4

Weed VCT suppressed root penetration of M. incognita consistently in both split-root trials (P ≤ 0.05), but vegetable VCT was only effective in Trial II (Fig. 5A). However, neither VCTs suppressed R. reniformis root penetration (P > 0.05, Fig. 5B) despite showing a trend of suppression compared to the water control.

Figure 5

No nematodes were found in the water control. There were more bacterivorous nematodes in weed vermicompost than vegetable vermicompost in Trial I (P ≤ 0.05, Fig. 6A), but more omnivorous nematodes were found in weed vermicompost than vegetable vermicompost in Trial II (P ≤ 0.05, Fig. 6B). No fungivorous, herbivorous or predatory nematodes were detected in either vermicompost examined.

Figure 6

Drenching of both types of VCT did not affect shoot, root, and pod weights of cowpea (P > 0.05, data not presented). However, weed VCT reduced root-gall index compared to the water control in both trials (P ≤ 0.05, Fig. 7). Both VCTs did not reduce the number of M. incognita and R. reniformis in the soil in neither of the trials (Table 2). In fact, the weed VCT treatment increased the abundance of Meloidogyne spp. in Trial II compared to the water control at the end of the experiment. Although abundance of bacterivores and fungivores were not affected by VCT drenching compared to the water control on all sampling dates in both field trials (P > 0.05), weed VCT increased omnivorous nematodes in Trial II by >5-fold at two months after cowpea planting (P ≤ 0.05, Table 2).

Figure 7