Entomopathogenic Nematode Control of the Asiatic Garden Beetle, Maladera castanea, in Corn

Objective 1. Screen local soils for presence of naturally occurring EPNs

Identification of isolated EPNs. Based on the resulting waxworm and AGB grub cadaver colors (Fig. 4) and confirmation of the presence of EPNs under the microscope with a basic assessment of relative size and body plan, three species, in addition a fungus, Metarhizium anisopliae (image not shown), were successfully isolated from every field sampled. The nematode species include (Fig. 4): Heterorhabditis bacteriophora (Hb), H. megidis (Hm), and Steinernema glaseri (Sg). M. anisolpliae (Ma) infections are observed when the insect host turns a greenish-gray color and white structures form on top of it that secrete droplets. Efforts are currently being made to confirm the identification of these species using the following protocol that was developed by Camila Hofman et al. (in press) to sequence the mitochondrial cytochrome oxidase subunit I (COI) gene.

Nematode Identification: mtCOI DNA barcoding. Single nematodes for each cadaver color were placed in 1.5 ml microcentrifuge tubes, then onto glass coverslips in 18 μl of sterile water. DNA extractions followed a modified protocol (Powers et al. 2014) where nematodes were sliced into pieces using a razor blade under a dissecting microscope. Mashed nematodes were stored in molecular grade water in individual 0.25 ml PCR reaction tubes until PCR reactions were performed. The primer set used to amplify the mitochondrial cytochrome oxidase subunit I (COI) gene consisted of COI-F1KF (29bp, 5’- CCTACTATGATTGGTGGTTTTGGTAATTG-3’) and COI-R2KF (23bp, 5’- GTAGCAGCAGTAAAATAAGCACG-3’) (Kanzaki and Futai 2002). Used. PCR amplification reactions will consist of 6.4 ml of ddH2O, 1.8 ml of each 20 mM primer, 15 ml of 2XJumpStart RED Taq ReadyMix (Sigma-Aldrich, Inc. St. Louis, MO) RED Taq ReadyMix (Sigma-Aldrich, Inc. St. Louis, MO) and 5 ml DNA template from the macerated nematode, for a total reaction volume of 30 ml. PCR cycling protocol will consist of a hot-start and an initial denaturation of 5 minutes at 94°C followed by 45 cycles of 15 seconds at 94°C (denaturation), 15 seconds at 55°C (annealing), 60 seconds at 72°C (extension) and a final extension step of 5 minutes at 72°C. All PCR will be conducted in a thermal cycler at the Ohio State University Department of Entomology. To confirm successful amplifications, 3 ml of PCR products will be loaded into 1% agarose gels stained with GelRed Nucleic Acid Gel Stain (Biotium, Hayward, CA, USA) in 1× Tris-acetate-EDTA (TAE) buffer. Gels will be placed into electrophoresis with 0.5X Tris-Borate-EDTA (TBE) running buffer for 35 minutes at 155V. DNA templates will be sequenced via bidirection Sanger sequencing at Functional Biosciences (Madison, WI), then edited and aligned on Geneious (Biomatters Inc., Newark, NJ). Excluding the primers, amplification products should yield 658-bp for resulting sequences. 

            Initial attempts at Sanger sequencing rendered a majority of bases as ambiguous and ultimately useless. Attempts will be made to clean up the PCR products and resequence them at the Ohio State University Molecular and Cellular Imaging Center in Wooster, OH. If that does not work, then individual nematodes will be macerated following the Powers et al. (2014) protocol verbatim and re-sequenced at the MCIC. Resulting sequences will be uploaded into Geneious Prime software and the consensus sequences formed, then be submitted to NCBI GenBank following publication of the resulting manuscript from this research. The sequences will be used to create a maximum likelihood phylogenetic estimation using MEGA X software following models selected by Bayesian Information Criterion scores. These results will increase our understanding of the genetic makeup of naturally occurring nematode populations which could give us insight to how similar they are; should species from nearby fields be genetically distinct, then a more targeted approach might be necessary when isolating local EPNs to mass rear and apply back into the system to ensure compatibility.

EPN spatio-temporal dynamics. The overall species composition of entomopathogens overall (Fig. 5A) and in 2018 (Fig. 5B) and 2019 (Fig. 5C). Three of the four sites sampled only in 2018, all of which had the lowest grub densities had at least 50% of positive samples contain Sg and no sampled containing Hm. The other sites were much more varied in species composition. All species were isolated from both the upper 5 cm and lower 15 cm of each soil core. Kaya et al. (1993) observed that Steinernema spp. occur near the soil surface and Heterorhabditis spp. are lower in the soil profile (Kaya et al. 1993). It is possible that these results are conflicting because the nematodes are responding to the presence of AGB grubs. In June, the grubs are found lower underground as they begin to enter pupation, while grubs in the fall are closer to the soil surface following harvest (personal observation).

          Overall there were significant differences in the number of samples containing each entomopathogen over the three evaluation periods (Fig. 7). In general, populations for Hb (Fig. 8A) and Ma (Fig. 9B) declined from September 2018 to June 2019, while Hm (Fig. 8B) increased and Sg (Fig. 9A) remained consistent. Northwest Ohio saw unprecedented rainfall events in Spring and early summer 2019 which left many fields inundated for days which extensively delayed and even prevented plantings for many farmers. Wauseon, OH saw at least 5” more rain in 2019 than the annual precipitation rate. During this time, the AGB adult population was only 2/3 that of what was observed in 2018. The excessive rains continued through the summer with July seeing more than 7” of rain. Ma (Fig. 9D) and Sg (Fig. 9C) populations remained constant through October 2019 while Hb (Fig. 8C) populations increased slightly and Hm (Fig. 8D) decreased. Hb (Fig. 8E) and Ma (Fig. 9F) were the only two entomopathogens to significantly decline from September 2018 to October 2019; the presence of Hm (Fig. 8F) and Sg (Fig. 9E) did not vary over time. These findings suggest that Hm does successful when soil moisture is high relative to the other entomopathogens, while Hb and Ma are highly susceptible to excessive soil moisture. Although soil moisture was not directly evaluated in this study, fields were observed to be oversaturated from May to July, with many fields supporting large ponds of water. If these environmental extremes become more routine, it may be necessary to select for species adapted to oversaturation, or drought, to ensure maximum efficacy and control.

            The combination of variables assessed in the multiple regression analysis elicited a significant and slightly positive relationship (R²=0.054) between these variables, particularly site (t=0.11, df=1, P=0.0204), evaluation period (t=-2.89, df=1, P=0.0041), and crop either being soybean or corn (t=2.34, df=1, P=0.0196), with the average number of grubs per plot (F=2.89, df=7,352, P=0.0060). There were no significant relationships between the number of grubs and presence of each different entomopathogen species (P>0.05). Ma, which was not included in the analysis since it behaved colinearly with other variables, declined so much following the excessive rains of 2019 that it was barely isolated from samples in June and October 2019. 

            Principal component analysis (Fig. 10) was used to determine whether there were any consistencies among plots with varying numbers of grubs, or site location site were structured by a combination of the presence of different entomopathogen species, sand, silt and clay content, evaluation period, and crop. Since silt, clay, and Meterhizium anisopliae were colinear with other variables in the multiple regression analysis, they were discarded. Prin1, or principle component 1, was heavily weighted by evaluation period and crop. Prin2, or principle component 2, is weighted most by sampling location and the presence of sand, and to a lesser extent by the presence of Heterorhabditis bacteriophora. In general, the sites on this figure show some homogeneity as they overlap to a certain extent, although some sites lean further in one direction or the other.

Future directions. The resulting sequences will provide positive controls for us to confirm additional isolated nematode species. We will continue to sample each field for entomopathogens by taking 20 composite samples per field in June 2020 and Fall 2020. I am currently learning how to use GIS to accompany these analyses. A manuscript will be prepared from these results and submitted to PLOS ONE following completion of this work.

Objective 2. Rear isolated EPN species using established “in-vivo” techniques

Each species was successfully reared following the protocol established by Testa and Shields (2017). Each species is currently being reared/preserved in several ways: populations were established in vegetative areas around the OARDC in Wooster, OH. Autoclaved soil was inoculated with each species and stored in vented containers in the dark at room temperature. Nematode colonies are being maintained in waxworms in the lab. In order to maintain genetic variation and field-like attributes, nematodes will be sampled in the spring and augmented with those previously collected as recommended by Dr. Elson Shields. These species will be mass reared using waxworm cups prior to field trials in the spring.

Objective 3. Evaluate AGB susceptibility to these nematodes in corn

Using a conservative rate of ~11,824 nematodes per ft², all four nematode treatments successfully infected and killed AGB, albeit at different rates (Fig. 11). (It is important to note that the number of grubs used in this study was a limitation as we are unable to rear the species in a controlled setting and we must drive 2-3 hours to collect grubs from the field. In addition, grubs are approximately the size of sand grains in the summer months, so we had to wait until mid-Fall to collect grubs that were a reasonable size to work with as our bait stations failed due to other animal species, which occurred around the time grubs were turning into adults). Hb and Hb + Hm treatments caused the highest mortality around 40% and were significantly different from the control which received no nematodes. Sg and Hm alone killed approximately 10% and 20% of grubs, respectively, but did not vary from the water control. Despite mortality rates below 50%, the rate used was only a fraction of what would be applied in a production setting, suggesting that actual field mortality rates may be much higher. Additional factors that may have  impacted nematode infection rates (e.g. soil type and grub instar) were compared in separate two-way ANOVAs using proc reg in SAS 9.4; grub infection rates did not vary for sandy and loamy soils or among 2^nd and 3^rd instar grubs (P>0.05). These results were largely consistent with those evaluating commercially available nematodes against AGB in turf grass systems. Unlike these studies, our study isolated nematodes from the local environment as these species will be better adapted to local environmental extremes such as drought and temperature extremes and should not require annual augmentation into the field; previous studies in upstate New York showed that locally isolated EPN can reduce pest populations in field cropping systems and persist for over 7 years after initial application.

Figure 12. Average percent of grubs that died from EPN treatments including a water control, 1000 IJs of either Hb, Hm, or Sg, or 500 IJs of Hb and Hm, respectively. This experiment was conducted in the greenhouse with V2 organic corn plants grown in 32 oz deli containers. Letters denote significant differences among treatments from multiple comparisons tests.

            Plant growth characteristics (root, shoot and total plant weights) were similar for plants of all treatments (Fig. 12). Interestingly, plant weights were numerically the smallest for Hb, one of the treatments that caused the highest AGB mortality. When looking at the roots and shoots of plants from this treatment, it was apparent that the grubs fed on the main roots closest to the kernel at a higher rate for this treatment than the others. This suggests there is likely a very low economic threshold level for AGB grubs in corn, thus necessitating the need for novel management tools like EPN, especially since insecticide seed treatments appear to be ineffective against this pest. These results were presented at the Entomological Society of America’s 2018 Annual Meeting in Vancouver, BC on November 12, 2018. I was recently invited by Paul Georgen from Pioneer Seed in Indiana to present these results to local farmers in north central Indiana on January 24, 2019. In addition, I will hold a workshop in 2021 to educate farmers in NW Ohio about EPN, how to mass produce them, and how to apply them using their own equipment. This workshop will accompany an annual Asiatic garden beetle meeting that I have held the past 2 years.

Although initially planned for the 2018 field season, the field evaluations will begin in 2020 using a microplot trial outdoors at the OARDC in Wooster, OH (Fig. 13). A grant extension was obtained due to a brief teaching appointment during Fall 2017 when this research was proposed to begin. In 2019, excessive rains drowned nearly 2/3 of the Asiatic garden beetle populations and prevented collaborators from planting until after the grubs pupated. The persistence and efficacy of EPN against AGB grubs (and other soil dwelling pests) will be evaluated in a microplot study in which microplots consisting of 12” tall PVC pipes with a 12” diameter placed in the ground will contain either sand or loam soil and receive a treatment of water, Hb, Hm, or Hb and Hm. A randomized complete block design with four replicates per treatment will be used for this study. Two organic corn seeds will be planted beginning of May and thinned following germination. Weeds and other pests will be managed per commercial recommendations. EPNs will be mass produced in the lab using containers of waxworms. They will be applied using a backpack sprayer that has the nozzle screens removed after plant. Depending on availability, at least 5 grubs and 5 waxworms will be placed in separate EPN bait stations in each microplot. The bait stations will be collected after 72 hours and any discolored grubs or waxworms will be placed on a White trap to isolate the EPNs for identification. Additionally, 3 soil cores will be taken and combined to create one composite sample per microplot in parallel with the baiting technique (Fig. 14). Every 3 months through Fall 2021 grubs (if able to collect) and waxworms will be placed in each microplot, and one composite sample taken, to monitor EPN persistence and efficacy over time. These results will be compiled into a manuscript in combination with the greenhouse trial to be submitted to the Journal of Economic Entomology following the completion of all work.