Can the botanical azadirachtin replace phased-out soil insecticides in suppressing the soil insect pest Diabrotica virgifera virgifera ?
| Main Authors: | Toepfer, S, Toth, S, Szalai, M |
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| Format: | info dataset Journal |
| Bahasa: | eng |
| Terbitan: |
, 2021
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| Subjects: | |
| Online Access: |
https://zenodo.org/record/4318642 |
Daftar Isi:
- Can the botanical azadirachtin replace phased-out soil insecticides in suppressing the soil insect pest Diabrotica virgifera virgifera ? Background Due to recent bans on the use of several soil insecticides and insecticidal seed coatings, soil-dwelling insect pests are increasingly difficult to manage. One example is the western corn rootworm (Diabrotica virgifera virgifera, Coleoptera: Chrysomelidae), a serious root-feeder of maize (Zea mays). We investigated whether the less problematic botanical azadirachtin, widely used against above-ground insects, could become an option for the control of this soil insect pest. Methods Artificial diet-based bioassays were implemented under standard laboratory conditions to establish lethal dose curves for the pest larvae. Then, potted-plant experiments were implemented in greenhouse to assess feasibility and efficacy of a novel granular formulation of azadirachtin under more natural conditions and in relation to standard insecticides. Results Bioassays in three repetitions revealed a 3-day LD50 of 22.3 μg azadirachtin per ml which corresponded to 0.45 μg per neonate of D. v. virgifera and a 5-day LD50 of 19.3 μg per ml or 0.39 μg per first to second instar larva. No sublethal effects were observed. The three greenhouse experiments revealed that the currently proposed standard dose of a granular formulation of 38 g azadirachtin per hectare for in-furrow application at sowing is not enough to control D. v. virgifera or to prevent root damage. At 10x standard-dose total pest control was achieved as well as the prevention of most root damage. This was better than the efficacy achieved by cypermethrin-based granules and comparable to tefluthrin- granules, or thiamethoxam seed coatings. The ED50 for suppressing larval populations were estimated at 92 g azadirachtin per ha, for preventing heavy root damage 52 g /ha and for preventing general root damage 220 g /ha. Conclusions There seems clear potential for the development of neem-based botanical soil insecticides for arable crops such as maize. They might become, if doses are increased and more soil insecticides phased out, a promising, safer solution as part of the integrated pest management toolkit against soil insects.
- One data file is related to artificial diet – based laboratory bioassays. One data file is related to potted -plant greenhouse experiments. Artificial diet – based laboratory bioassays Experimental setup To assess lethal doses of azadirachtin on neonates of D. v. virgifera, artificial diet-based bioassays with different dosage were conducted against neonates in three replicates under controlled semi-sterile conditions (Tab.1). Azadirachtin from a common fluid formulation was compared with a novel granular formulation. The insecticides cypermethrin, tefluthrin and imidacloprid served as positive control. Sterilised tap water or no treatment at all served as negative controls. Each bioassay consisted of 3 to 6 polystyrene plates of 96 wells each (07-6096 of Biologix Ltd., USA, or Costar 3917 of Corning Inc., USA). Each well was of 330 μl volume with 5 mm in diameter and 10 mm height, and had a 0.34 cm2 surface. Each treatment was applied to 8 wells of each plate per bioassay. Each treatment-dose combination was tested in at least in two true replicates. The larval diet for a bioassay had been prepared one day before treatment and infestation. The diet was prepared under semi-sterile conditions following methods of Sutter et al. (1971); Pleau et al.(2002), Moar et al.(2017); and P Clark, E. Boland (2016, Genective, pers. comm.). A commercial southern corn rootworm diet was used (Frontier #F9800B, Frontier Scientific Ltd., USA), but maize and food colour were added. This diet consists of D(+) sucrose, vitamin-free casein, cellulose, Wesson's salt mix, methyl paraben fungicide, sorbic acid, cholesterol, raw wheat germ, Vanderzant's vitamin mix, raw linseed oil, streptomycin sulphate antibiotic, and chlortetracycline antibiotic. For 100 ml of diet, 13.8 g of the #F9800B diet was grinded and added to 88 ml fluid 60 to 70 °C agar (1.5 g agar CAS 9002-18-0, Chejeter, Japan in deionized water). After blending and cooling to 55 to 60 C, 0.75 g grinded lyophilized maize roots were added (GLH5939 Pioneer, USA, or Phileaxx RAGT, Hungary) as well as 0.1 g green food colour for better larvae observation (Les Artistes, France) . Then, 1.7 to 1.8 ml 10%w/v KOH were added to reach a pH between 6.2 and 6.5. This mix was blended again, and then stirred at 50 to 55 °C. Then, 190 μl diet was pipetted into each 330 μl well filling each to around 2/3rd (repeater pipette P-8, Topscien Co., Ltd, China). Then, plates with diet were allowed to dry in a laminar flow cabinet during 45 minutes, and then stored at 3 to 5 °C overnight. The following day, treatments were applied. This is, 20 μl of a treatment were applied to the 0.34 cm2 diet surface in each well (10 to 100 μl pipette, Biohit Proline, Finland). Order of treatments were shifted every other plate to avoid edge effects. Then, plates were dried for 1 to 1.5 hours, and then cooled for 1 hour in a 3 to 5 °C fridge. Two weeks prior the bioassays, soil dishes with freshly laid eggs had been removed from D. v. virgifera adult rearing cages to allow sufficient incubation time until egg hatch. Eggs were washed with cool tap water with <0.01% NaOCl through a 300 μm mesh sieve. Around 5000 eggs were transferred to sterilise, slightly moist river sand (< 200 μm grains) in Petri dishes. They were incubated at 24 ± 2 °C in darkness for 8 to 12 days until hatching started. One day before a bioassay, the ready-to-hatch eggs were again washed and sieved. Eggs were then again placed on sterile moist sand onto slightly moist tissue paper into a dish to allow clean hatching conditions of new neonates and their use for bioassays. One neonate larva was placed per well using a fine artist brush. A fast-moving, healthy-looking larva was chosen, and lifted from the end of abdomen with the brush, moved towards a well surface, and allowed to crawl off the brush onto the diet. Larvae were not placed in treatment column order but rectangular to avoid systemic errors. After every 12th larva, the brush was cleaned in 70% ethanol followed by sterile tap water. The filled plate was closed with an optically clear adhesive qPCR seal sheet (#AB-1170, Thermo Scientific, USA or #BS3017000, Bioleader, USA) allowing data assessments without opening the plate. Four to five holes were made with flamed 00-insect pins into the seal per well to allow aeration. The plates were incubated at 24 ± 2 °C and 50 to 70% r.h. in dark in a ventilated incubator (Friocell 22, MMM Medcenter, Munich, Germany) for 5 days. Data assessments and analyses After 3 and 5 days of incubation, larval mortality, stunting, feeding and contamination were visually assessed through the clear seals using a stereomicroscope (10x magnification, SMZ-B4, Optec, Chongqing, China). Data from a plate were only used when the natural mortality threshold in the untreated control had not been reached, i.e. no more than 3 dead per 8 larvae per column of wells (37.5% threshold). This is in contrast to common practices in bioassays with other insects where the quality acceptance is usually <10% natural background mortality (Dulmage et al., 1990). However, this is rarely achievable with rootworm larvae as the artificial diets known to date remain suboptimal (B. Hibbard, University of Missouri, 2019, pers. comm.; Huynh et al. 2018)). Stunting was qualitatively assessed as an indicator for sublethal effects in comparison to the size and form of larvae in the untreated control. Feeding was assessed through observing food remains, frass, and diet in the larval gut to assure that diet and a treatment had been ingested. The coefficient of variation (CV) was determined in each bioassay as a measure of data precisions. A CV should ideally be < 0.2, and at a CV of > 2 further bioassays would be needed(Dulmage et al., 1990). In our experiments, the CVs of 1.2 for bioassay 1, 0.4 for bioassay 2, 0.7 for bioassay 3, and 0.8 for all bioassays, indicated good quality of data (Table in additional file). Larval data were compared between treatments within each experiment using Chi-Square statistics (because of nominal data type) with an fdr-correction of p-values (Benjamini and Hochberg 1995). To allow across-experiments comparisons, data were standardised to the untreated control data. Distributions of data were investigated using histograms as well as normal and detrended normal probability Q - Q plots (Kinnear and Gray 2000). Equality of variances was assessed using Levene`s test. When data appeared normal distributed, influences of treatments were analysed through unifactorial GLM and multiple comparisons were applied using Tukey HSD post hoc comparison of data of equal variances and Games Howell post hoc comparisons for unequal variances. Logistic regression analyses were applied to assess the dose response of each treatment including lethal dose leading to 50% or 90% mortality (LD50/90) (R packages MASS and DescTools (R Development Core Team 2020). Potted -plant greenhouse experiments Experimental setup To assess the efficacy of azadirachtin granules against D. v. virgifera larvae under semi-natural conditions, three systematic controlled trials were conducted using infested potted - maize plants in a greenhouse. As positive control served tefluthrin fine granules, cypermethrin microgranules and thiamethoxam - seed coating (Tab. 2). As negative controls served untreated infested plants as well as untreated uninfested plants. Each treatment was applied into the soil of three to four systematically arranged blocks (= replicates) of five pots. This totalled 15 to 20 data points (= sample size) per treatment per experiment. In detail, each pot (plastic garden pot, 15 cm inner diameter x 10 cm height, 2 litres) was first filled with 1 litre sterilised soil. Then, two maize seeds (hybrid Szegedi 386, GK Hungary in experiment 1 and 3 , or Futurixx, RAGT, France in experiment 2). Then, 200 ml water were applied to each pot. Treatments were applied either as granules along a 2 cm wide strip across the 10 cm diameter of the pots, or as seed coating. Then, 1/2 litre soil was added burring the treatment and seed 3 cm into the soil leading to a soil surface of 14 cm diameter. The used soil contained 77%m/m sand, 8% loam, 15% clay, 2.8 % humus, 1.7 % CaCO3, 0.1% salts, and had a Ph of 7.7 (analysed by Szolnoki Talajvedelmi Laboratorium, Hungary). It had a soil bulk density 0.9 to 1.1 g cm-3 and a 7 to 11 % soil moisture (w% = grav. %). A temperature of 20 ± 5 °C and a relative humidity of 97 ± 3 % were recorded 5 cm deep in the soil in the pots as well as 24 ± 4 °C and 44 ± 13 % in the air 1 m above the pots using climate data loggers (PeakTech 5185 data logger, Germany). Plants germinated between 4 to 12 days after sowing. Maize pots were infested with 50 viable ready-to-hatch eggs per plant in experiment 1 and 3 or with 100 eggs in experiment 2. At this point in time, the majority of plants was at 3 leaf stage (height 15 to 20 cm). The eggs were applied in 0.15 to 0.2 % aqueous agar with a standard pipette (1 to 5 ml, Eppendorf AG, Germany) in half-portions into two 50 mm deep holes 20 to 30 mm distant from both sides of the plant. A portion of eggs was incubated on moist filter paper at 20 °C in the laboratory to estimate emergence patterns (5 dishes with 10 to 20 eggs each per experiment). They revealed an emergence start 9 ± 5 days after placement. Hatching duration was 10 ± 4 days. Hatching rate was 47 ± 30, 47 ± 23 % and 56 ± 19, leading to 24 hatched larvae per pot in experiment 1, 47 larvae in experiment 2, and 28 larvae in experiment 3, respectively. This indicates a medium, but consistent egg quality across experiments, and is comparable to similar studies of Xie et al. (1991). Data assessments and analyses Selectivity of the test agents was assessed by recording germination rate, plant phenology and phytotoxicity. Leaf number, plant height and the BBCH growth stage were assessed weekly as well as phytotoxicity according to Anonymous (2009). At the expected second and early third instar stage, numbers of surviving larvae, root damage and above-ground biomass were assessed. This was 52 ± 18 days after planting and treatment, thus 40 ± 7 days after infestation. Each maize plant was pulled out of the soil, and gently shaken to remove loosely adhering soil particles from roots. Each maize plant was cut 1 cm above roots, and fresh weight, leaf number and plant height were measured. Then, the soil and root of each pot was placed onto a plastic screen for drying out and letting surviving larvae exit and drop onto the wet tissue paper in a tray below following a Berlese approach (Dent and Walton 1998). Larvae and their instars were counted 1, 3, 5 and 7 days later. The untreated control was aimed to have a minimum level of infestation with 2nd or 3rd instar larvae of 20% to validate the results on agent efficacies. In all experiments, more than 90% of pots of the infested untreated control yielded larvae. The infested control lead to 6 ± 5 second or third instar larvae per 100 applied eggs. One day after Berlese-placement, the dried roots were removed, gently shaken to remove remaining soil, soaked in water for 5 minutes, and then washed in 1% NaOCl and then water for one minute to allow the assessment of root damage. Damage was rated using two scales recommended by EPPO (Anonymous 1999); this is, (1) the non-linear 1.0 to 6.0 traditional IOWA scale (Hills and Peters 1971) which slightly overestimates minor damage; and (2) the linear 0.00 to 3.00 node injury scale (Oleson et al. 2005) which measures only destroyed roots and therefore misses minor damage. To avoid subjective bias on these ratings, root damage was estimated independently by the experimenters, neither of whom knew whether the roots were from a treated or untreated pot. Distributions of data were investigated using histograms as well as normal and detrended normal probability Q - Q plots (Kinnear and Gray 2000). Equality of variances was assessed using Levene`s test. Influences of treatments on assessed factors were analysed through GLM analyses or through independent samples Kruskal-Wallis H test. Tukey HSD post hoc multiple comparison tests were applied following GLM in case of equal variances, and Games Howell test in case of unequal variances. Logistic regression analyses were applied to assess the dose response of each treatment including the effective dose leading to 50% suppression of the larval populations or root damage prevention (ED50) The mean corrected efficacy of each treatment was calculated relative to the untreated control, this is corrected efficacy % = 100 × (larvae or damage in control plots – larvae or damage in treated plots)/maximum (larvae or damage in control or treated plots) (Toth et al. 2020). As the 1.0 to 6.0 IOWA root damage scale is a non-linear ordinal scale, and a value of 1 equals no damage, the damage data were converted to a 0.0 to 5.0 scale to estimate percent damage prevention across experiments. Results from azadirachtin treatments were validated in relation to the results from the corresponding positive controls of standard insecticides