DOI: 10.15482/usda.adc/27982133.v1
title: Data from: Non-consumptive effects of parasitoids and predators in stored products: The case of <i>Theocolax elegans </i>and other field-collected predators on the foraging of lesser grain borer and rice weevil

description: <b>Insects</b>Beetles used in this study were obtained from stock colonies maintained at the USDA Agricultural Research Service’s (ARS) Center for Grain and Animal Health Research (CGAHR) in Manhattan, KS, USA. Colonies of <i>R. dominica </i>and <i>S. oryzae</i> were reared on organic whole wheat kernels that had been tempered to 15% grain moisture. To subculture, a total of 50 adult individuals were placed on 200 mL of grain in a mason jar (capacity: 473 mL) and given 14 d to mate and lay eggs. At the end of that period, adult hosts were removed by sieving with a #10 sieve (2.00 mm; W.S Tyler Inc., Mentor, Ohio), and colonies were allowed to age for 3-weeks prior to using beetles as hosts for parasitoid rearing. <i>Theocolax elegans </i>were maintained separately on two different hosts, either <i>R. dominica </i>or <i>S. oryzae </i>for at least three full generations. Freshly emerged, healthy <i>T. elegans </i>were used for the experiments below. All colonies of parasitoids were maintained in a separate environmental chamber than host-only colonies to prevent cross-contamination. Colonies were maintained in mason jars and stored in an environmental chamber under constant conditions (27.5°C, 60% RH, 14:10 L:D).<b>Interactions with Predators</b>Laboratory studies were performed in 2022 and 2023 at the USDA Center for Grain and Animal Health Research (Manhattan, KS, USA). From July–October of each year, predators were collected weekly from local post-harvest food facilities, including the Kansas State Agronomy Farm (GPS: 39.2062227, -96.5951959), where <i>S. oryzae </i>and other stored product pests are abundantly found (Morrison et al. 2025[1] ). Most predators used in trials were collected by sweep netting (Bioquip Products, Inc., Rancho Dominguez, CA) sampling vegetation adjacent to grain bins or by hand collection and held temporarily in 1-gal (=3.98 L) Ziplocks, then immediately brought back to the lab in a cooler on insulated ice packs. In the lab, insects were processed by individually placing predators into a 950-mL mason jar with 10 <i>S. oryzae </i>from colonies. The predators were identified to family (Marshall 2006, Paquin et al. 2017). Mason jars with predators and <i>S. oryzae </i>were then placed on shelves in an environmental chamber set to constant conditions (27.5°C, 60% RH, 14:10 L:D). After 24 h, the jars were checked, and the number of <i>S. oryzae </i>consumed was recorded as well as the presence of any self-aggregation behavior of <i>S. oryzae </i>together and away from the predator, which was taken to be evidence for non-consumptive effects in the presence of the predator. The results of predators were only included when there were n = 3 or greater number of replicates.<b>Ethovision</b>Video-tracking coupled with Ethovision software v.14.0 (Noldus, Inc., Leesburg, VA: Noldus et al. 2002) was used to investigate the impact of natural enemy kairomones on the mobility and orientation of <i>R. dominica </i>and<i> S. oryzae </i>over short distances. This system has previously been used for analyzing the mobility and foraging behaviors of stored product insects (Wilkins et al. 2020; Ponce et al. 2022). Six arenas consisting of Petri dishes (VWR Petri dishes, 100 × 15 mm) with an 85-mm filter paper (Grade 1, Whatman, GE Healthcare, Chicago, IL) adhered to the bottom using double-sided sticky tape were arranged 80 cm below a network video camera (GigE, Basler AG, Ahrensburg, Germany). The movement of individual insects within each arena was simultaneously recorded on an adjacent computer. Four zones were monitored in Ethovision, including the two halves of the Petri dish (i.e. treatment half vs control half) and two 1 cm diameter zones nested in the middle of each half where stimuli were applied (treatment stimulus zone and control stimulus zone). The position of treatments was randomized between replicates and a total of n = 12 replicate assays were conducted for each treatment. For each assay, a single insect was introduced into the center of an arena and its movement was tracked for a total of 10 min. Several measurements were summarized in Ethovision including cumulative distance moved (cm), instantaneous velocity (cm/s), frequency of entering each stimulus zone, latency to enter each stimulus zone, cumulative time in each stimulus zone, and cumulative time in each half of the arena.<b>Parasitoid cues</b>Both headspace extracts (wasp extract experiments) and adult wasps (adult wasp experiments) were employed as stimuli in experiments examining influence of parasitoid cues on <i>R. dominica </i>and <i>S. oryzae </i>movement and foraging. Treatment stimuli employed for wasp extract experiments consisted of clean solvent (control), headspace extract collected from 100 g of uninfested wheat (wheat), and headspace extract collected from colonies of conspecifics parasitized by <i>T. elegans</i> (parasitoid) applied in 10 μL aliquots to 10 mm filter paper discs. Prior to addition of filter paper to arenas, the solvent was allowed to evaporate for 15 s. Adult wasp experiments employed a single wheat kernel (wheat), two female <i>T. elegans </i>(parasitoid), and no stimulus (control) as treatment stimuli. Ethovision experiments were blocked by presence of wasps to account for potential spillover wasp odors to neighboring arenas. Control zones either lacked stimuli (adult wasp experiments) or contained a 10 mm filter paper disc to which 10 μl of clean dichloromethane solvent was applied (wasp extract experiments).<b>Predator cues</b>As Orthoptera were most commonly encountered in field plots and consumed <i>S. oryzae </i>at relatively high rates, experiments examining the response of <i>S. oryzae </i>and <i>R. dominica </i>to predator cues focused on one orthopteran family, namely Gryllidae. <i>Gryllus pennsylvanicus </i>(hereafter,<i> </i>gryllids) were collected from the Kansas State University Agronomy Farm by deploying 9 bottle traps (5 × 10 cm D:H)[4] flush with the ground spaced 5 m apart among grain bins with <i>S. oryzae</i> documented in the area. The pitfall traps were 3D-printed (Lulzbot Taz 6) and baited with 5 g of cornmeal. Traps were checked daily during August–September 2024. Two experiments were conducted to assess the impact of gryllid cues on <i>R. dominica </i>and <i>S. oryzae </i>movement. The first experiment examined the impact of olfactory stimuli on <i>R. dominica </i>and <i>S. oryzae </i>movement, with treatment stimuli consisting of clean solvent and cricket headspace extracts. The cricket headspace extracts were prepared according to the headspace collections below. The second experiment examined the impact of predator visual cues with and without associated olfactory cues on <i>R. dominica </i>and <i>S. oryzae </i>movement. For this experiment, treatment stimuli consisted of clean solvent, clean solvent with visual cues, and cricket headspace extract with visual cues. Visual cues consisted of small (30 × 8 × 5 mm L:W:H) cricket models (Toyvian) that were first baked off at 75℃ for 30 min to ensure they were chemically inert.<b>Headspace Volatile Collections</b>To determine the role of chemical cues in mediating nonconsumptive interactions between stored product pests and their natural enemies, headspace volatiles were collected from predators (5 crickets, Gryllidae), colonies of <i>S. oryzae </i>and<i> R. dominica </i>that were parasitized by <i>T. elegans</i>, and uninfested wheat kernels. All samples were collected using a headspace collection system (after Van Winkle et al. 2022). An activated carbon filter was employed to remove background volatiles from central air, which was then split between eight lines. Each piece of the system was connected using chemically inert PTFE tubing and fittings. Inline flowmeters (Volatile Collection Systems, Gainesville, FL) were employed to maintain a flow rate of 1 L/min through each line. Volatiles were collected on traps consisting of a drip tip borosilicate glass tube containing 20 mg of Poropak-Q absorbent between a stainless-steel screen (No. 316), borosilicate glass wool, and a PTFE compression seal (Volatile Collection Systems, Gainesville, FL). Volatiles were collected for 24 h, after which volatiles were eluted by pushing 150 µL of HPLC-grade dichloromethane (Sigma-Aldrich, St. Louis, MO) through the traps with N<sub>2</sub> gas. The eluent was collected in 2 mL screw-cap GC vials (Item#5191-8121, Agilent Inc., Santa Clara, CA, USA) with 150-μL glass inserts with polymer feet (Item#5181-8872, Agilent Inc.). All samples were magnetic capped with PTFE-backed silicone septa (Item#XXX, Agilent Inc.), sealed with PTFE tape, and stored at -20°C prior to use in behavioral assays and chemical analysis.<b>Chemical Analysis</b>Headspace volatile samples of 50 µL aliquots of each sample were transferred to new GC vials with 150-µL inserts for analysis by GC-MS. Prior to chemical analysis, 190.5 ng of tetradecane was added to each sample as an internal standard. Sample extracts were then run on an Agilent 7890B gas chromatograph (GC) equipped with an Agilent Durabond HP-5 column (30 m length, 0.250 mm diameter and 0.25 µm film thickness) with He as the carrier gas at a constant 1.2 mL/min flow and 40 cm/s velocity, which was coupled with a single-quadrupole Agilent 5977B mass spectrometer (MS). The split/splitless inlet was operated in splitless mode and maintained at 250°C during injection. The initial oven temperature of 40°C was maintained for 3 min, before increasing to 280°C at a rate of 10°C/min, where it was held for 3 min. After a solvent delay of 5.5 min, mass ranges between 35 and 550 atomic mass units were scanned. A mixture containing C8-C20 alkanes was employed to calculate Kovats index for all peaks. Preliminary identification of peaks was achieved by comparing spectral data and Kovats index with references in the NIST 14 library. Data was compiled using Masshunter Unknowns Analysis (Agilent Inc., Santa Clara, CA, USA) and compounds were aligned with the R package <i>uafR </i>(Stratton et al., 2024).