We compared all combinations of three commercial traps and five different attractants on the capture of stored-product insects for two consecutive years in three food processing facilities in Central Greece. Specifically, Facility 1 and 2 were pasta factories and Facility 3 was a flour mill. The traps that were used in the experiments were Dome Trap (Trécé Inc., USA), Wall Trap (Trécé Inc., USA) and Box Trap (Insects Limited, Ltd., USA). The attractants that were evaluated were 0.13 g of : 1) of PantryPatrol gel (Insects Limited, Inc., USA), 2) Storgard kairomone food attractant oil (Trece Inc.), 3) wheat germ (Honeyville, USA), 4) Dermestid tablet attractant (Insects Limited Inc., USA). The traps were inspected approximately every 15 days and rotated clockwise. The captured insects were transferred to the Laboratory of Entomology and Agricultural Zoology (LEAZ) at University of Thessaly for identification. The results indicated that there was a wide range of species within the three facilities throughout the trapping period, with the Indian meal moth, Plodia interpunctella (Hübner), the red flour beetle, Tribolium castaneum (Herbst) and the cigarette beetle, Lasioderma serricorne (F.), being the most abundant. Although there were noticeable differences among the different traps and attractants for the capture of certain species, all combinations provided comparable population fluctuation patterns. In general, Dome traps, baited with either the oil or the gel, were found to be the most effective. There are not much data available so far for the simultaneous comparable use of different trapping devices and different attractants in commercial facilities for long-term monitoring. Certain lures are marketed toward particular pests or classes of pests, while others might be more generic, multi-species lures. To shed light on this issue, we evaluated a series of combinations of floor traps and attractants, in three commercial facilities in Greece, for a period of two years. Our questions included both which trap was broadly most effective as well as whether different combinations of traps and types of attractants were delivering novel information about the stored product insect community. The traps include two types of floor traps, and the wall trap used in the USDA khapra beetle detection programs. The lures included the Insects Limited ™ dermestid tab that is more specifically focused on food kairomones for only that taxon, and the same company’s PantryPatrol gel, which uses wheat kairomones and the pheromones of multiple species, including dermestids. We also use the Trécé Storgard kairomone oil, and simple wheat germ, which are both multi-species kairomones with no pheromones. Resources in this dataset: Resource Title: 2018 and 2019 field trapping data File Name: kb_greek_data_ag_data_commons.csv Resource Description: 2.1 Storage facilities The storage facilities in which this study took place are located in Central Greece. The selection of these facilities was based on their size, the accessibility from University of Thessaly (UTH) personnel and the known historical presence of stored product insect species and other arthropods. The sampling was conducted in three types of storage facilities refereed as Facility 1, Facility 2 and Facility 3. Facilities 1 and 2 are pasta factories, with substantial quantities of soft and hard wheat, flour and bran, but also some barley and maize, while Facility 3 is a flour mill, mostly focused on soft wheat processing. The deployment of the traps on each facility was conducted at 18 June 2018, 4 July 2018, and 3 July 2018 for Facility 1, 2 and 3, respectively. 2.2. Traps, attractants and inspection The trap types that were used in our experiments were Dome Trap (Trécé Inc., USA), Wall Trap (Trécé Inc., USA) and Box Trap (Insects Limited, Ltd., USA). These traps have been proven effective for monitoring purposes based on previous studies (Toews et al., 2009; Athanassiou and Arthur, 2018; Gerken and Campbell, 2021). Four attractants (noted also as lures) were used in our experiments, which were 0.13 g: 1) PantryPatrol gel (gel, Insects Limited, Inc., USA), 2) Storgard™ Oil kairomone food attractant (oil, Trécé Inc.), 3) wheat germ (WG, Honeyville, USA), 4) Dermestid tablet attractant (bait, Insects Limited Inc., USA). Also, an additional series of traps was used without any attractant, and served as “control” (e.g., ctrl). In Facility 1, the different traps and attractant combinations were replicated two times. In Facilities 2 and 3, the combinations were replicated three times, based on larger space availability. For each Facility, the traps were inspected approx. every 15 days, with the exception of some intervals where access to the trapping areas was not possible (e.g. due to fumigations in certain areas etc.). The traps were rotated clockwise after each inspection. The attractants were replaced at 15-day intervals, while the traps were replaced whenever it was considered necessary (damaged or lost traps). All captured insects were transferred to the Laboratory of Entomology and Agricultural Zoology (LEAZ), Department of Agriculture, Crop Protection and Rural Environment, University of Thessaly. 2.3 Identification The morphological identification of the captured individuals was carried out up to the species level, or lowest taxonomic unit, whenever this was possible using taxonomic keys, but in general many specimens are referred to as taxa. The insects found were classified into species (species identification) using different taxonomic keys (Bousquet, 1990; Peacock, 1993; USDA 1991). Data dictionary: rfb = red flour beetle cfb = confused flour beetle hfb = hairy fungus beetle lgb = lesser grain borer stgb = saw-toothed grain beetle cb = cigarette beetle rw = rice weevil gw = granary weevil imm = indianmeal moth rgb = rusty grain beetle trogoderma = dermestid genus

Four to six-week-old larvae of Trogoderma variabile and Trogoderma inclusum were used for the experiment. Both strains were originally obtained from the field in north-central Kansas in 2016 and 2012, respectively. Colonies of these species were reared under controlled conditions in an environmental chamber set to a temperature of 27.5 °C, 65% RH, and 14:10 (L:D) h photoperiod. Both species were fed 300 g of ground dog food (SmartBlend, Lamb flavor, PurinaOne, St. Louis, MO, USA) with oats sprinkled on top and a moistened, crumpled paper towel placed on the surface in a 950-ml mason jar. Treatments The long-lasting insecticide-incorporated polyethylene netting (2 × 2 mm mesh, D-Terrance, Vestergaard Inc., Lausanne, Switzerland) included 0.4% deltamethrin, or control netting that was identical in physical properties but without insecticide. These were used with the movement assay. We assessed the movement in the vicinity of important pheromonal and food kairomones after exposure to LLIN or control netting. Food consisted of 0.01 g of organic, unbleached flour (Heartland Mills, Marienthal, KS, USA), and pheromonal stimuli included a broad spectrum, multi-species lure (PTL lure, IL-108-10, Batch#1288200321, Insects Limited, Westfield, IN, USA), including Trogoderma spp pheromone (Ranabhat et al. 2023a). In each replicate, we used a single pellet (white color), and affixed it in place so it did not move in a Petri dish using a 1 × 1 mm square of parafilm. For each replication of testing, we used a fresh lure. Movement Assay The movement of larvae after exposure to the 0.4 % deltamethrin LLIN or a control netting in response to food cues (using 0.01 g of flour) or with conspecific sex pheromones (using a single bead from a disaggregated PTL lure held in place with a small square of parafilm), was tracked in six individual arenas (100 × 15 mm D: H) with a piece of filter paper (85 mm D, Ahlstrom-Munksjö, Helsinki, Finland) lining the bottom for 30 min using a network camera (GigE, Basler AG, Ahrenburg, Germany) affixed 76 cm above and centered over the dishes. The Petri dishes were backlit using a LED light box (42 × 30 cm W:L LPB3, Litup, Shenzhen, China) to increase contrast and affixed in place with white foam board. The video was streamed to a computer and processed in Ethovision (v.14.5 Noldus Inc., Leesburg, VA, USA). Prior to use in the movement assay, larvae of T. variabile or T. inclusum were exposed to the 0.4% deltamethrin LLIN or a control netting for 1 min in a 21 × 21 cm square Petri dish, then their movement was tracked individually after a post-exposure holding duration of 1 min or 24 h. A small 1.1 cm hidden stimulus zone encircled each stimulus, midway and centered on each half of the arena wherein movement was tracked separately from each half of the arena (control vs. treatment). The total distance moved (cm), instantaneous velocity (cm/s), frequency of entering each half of the petri dish and stimulus zone, cumulative duration spent in each zone (s), and latency of entering each zone (s) over a 30 min trial period was logged after exposure to a given treatment. The control side of the arena remained empty. A total of n = 16 replicates were run per treatment combination for both species No-Choice Release-Recapture Assay A release- recapture experiment was conducted for the larvae of both T. variabile and T. inclusum where larvae were exposed to the 0.4% deltamethrin LLIN and control netting for 1 min. After exposure, treated insects were released at one corner of the sanded plastic bin (60 × 41.6 × 16.5 cm L:W:H ). A commercial pitfall trap (Dome Trap™, Trécé, Inc., Adair, OK, USA) that contained a PTL lure (used only white beads as above), or 0.01 g flour, or no stimuli (unbaited for control), was deployed in the opposite corner, diagonally across from the release point in the bin. The bins were located in a large (4.8 × 2.1 × 6 m, L:W:H) walk-in environmental chamber (Percival Instruments, Dallas County, IA, USA) set at constant conditions (27.5°C, 60% RH, and 14:10 L:D). A total of 10 larvae were released in each bin during each replicate. Treated larvae were given 24 h to disperse to the semiochemicals in each trap, and then the number of insects captured inside the trap, found on the bottom of the trap, on the stimulus half of container or on the non-stimulus half of the container were recorded. A total of n = 12 replicates were performed per treatment combination for the larvae of each species. Resources in this dataset: Resource Title: Ethovision Movement Assay File Name: ranabhat_etal_larval_dermestid_et_LLIN_olfactory_agdata_commons.csv Resource Title: No-Choice Release-Recapture Assay for Larger Cabinet Beetle File Name: ranabhat_etal_larval_dermestid_rr_lcb_LLIN_agdata_commons.csv Resource Title: No-Choice Release-Recapture Assay for Warehouse Beetle File Name: ranabhat_etal_larval_dermestid_rr_whb_LLIN_agdata_commons.csv

Two grain surface treatment insecticides (deltamethrin and pirimiphos-methyl were evaluated in laboratory assays as a surface treatment for maize to control adult Prostephanus truncatus and Sitophilus zeamais. Both insecticides were applied to 20 g of maize placed in a vial or to the upper one half, one fourth, or one-eighth layer of the maize. Insects were either added to the vials before or after the maize. Mortality, progeny production, and insect damaged kernels (IDK) were then evaluated for each vial. Introduction method (before or after) did not have any impact on any of the variables. Mortality was nearly 100% for all treatments for both insecticides for P. truncatus. Subsequently, progeny production and the number of insect damaged kernels was very low or zero for P. truncatus. Mortality for S. zeamais remained low across layer treatments for deltamethrin. However, S. zeamais was easily controlled by primiphos-methyl. The results of this laboratory study show that while deltamethrin and pirimiphos-methyl has some effectiveness as a layer treatment on a column of maize, efficacy will be dependent on the target species, and the depth of the treated layer, as well as the location on which the insects are present. Resources in this dataset: Resource Title: Grain Layer Experiment with P. truncatus & Sitophilus zeamais File Name: quellhorst_etal_layer_experiment.csv Description: Insect Mortality on Treated Maize and Progeny Production. For each replicate, 500 g of maize were treated with each insecticide or H2O (e.g., control) as described above. Before proceeding with the experiments, the grain moisture content (m.c.) was assessed, using a moisture meter (mini GAC plus, Dickey-John Europe S.A.S., Colombes, France). The standard plastic cylindrical vials of the Laboratory of Entomology and Agricultural Zoology (LEAZ) were used (3 × 8 cm in diameter by height, Rotilabo Sample tins Snap on lid, Carl Roth, Germany). These were filled with 20g of maize. In each vial, we treated either all the grain (1/1), 1/2, 1/4 or 1/8 of the maize with one of the two insecticides (deltamethrin or pirimiphos-methyl) at the labeled rate. We also either placed the insects at the bottom of the vial (before the maize has been added) or at the top (after the maize has been added). Sets A, B, and C were treated with insecticide on separate days. Insects were given 14 days before mortality counts were performed. After this interval, the mortality was assessed. It is difficult to estimate the upper 1/8 etc. of maize, therefore we based our experiments on ratios of 20 g treated, 20 g untreated, 10 g treated with 10 g untreated, 5 g treated with 15 g untreated and 2 g treated with 18 g untreated. The exact quantities of the samples were weighed with a Precisa XB3200D compact balance (Alpha Analytical Instruments, Gerakas, Greece). The upper rings of the vials were treated with Fluon (Northern Products Inc., Woonsocket, USA) to prevent insects from moving away from the grain and or escaping. The top of each vial also had small holes punched to allow ventilation. Each vial then received 10 P. truncatus adults of mixed sex and age from the Tanzania strain or 10 S. zeamais from Brazil. The vials were placed inside incubators set at 30°C and 65% R.H. After the parental mortality count, all adults were removed, and the vials with maize were returned to the incubator at the conditions indicated above. Sixty days later, the vials were opened again to check progeny production and the number of insect damaged kernels (IDK). For each combination, e.g., insecticide × insect species, there were three replicates with three subreplicates (total 3 × 3 = 9 vials or replicates per combination). There were 2 insecticides × 2 insect species × 4 grain treatments (1/1, 1/2, 1/4, 1/8) × 2 insect introduction methods (before or after) × 9 replicates/subreplicates = 288 vials total, 5760 g of maize, 10 insects per vial × 288 = 2880 total (1440 per LAGB and MW). We also had a separate set of vials for the control with no insecticide= 9 × 2 insect species = 18 vials, 360 g of maize, and 180 insects (90 per species).

Our goals with this dataset were to 1) isolate, culture, and identify two fungal life stages of Aspergillus flavus, 2) characterize the volatile emissions from grain inoculated by each fungal morphotype, and 3) understand how microbially-produced volatile organic compounds (MVOCs) from each fungal morphotype affect foraging, attraction, and preference by S. oryzae. This dataset includes that derived from headspace collection coupled with GC-MS, where we found the sexual life stage of A. flavus had the most unique emissions of MVOCs compared to the other semiochemical treatments. This translated to a higher arrestment with kernels containing grain with the A. flavus sexual life stage, as well as a higher cumulative time spent in those zones by S. oryzae in a video-tracking assay in comparison to the asexual life stage. While fungal cues were important for foraging at close-range, the release-recapture assay indicated that grain volatiles were more important for attraction at longer distances. There was no significant preference between grain and MVOCs in a four-way olfactometer, but methodological limitations in this assay prevent broad interpretation. Overall, this study enhances our understanding of how fungal cues affect the foraging ecology of a primary stored product insect. In the assays described herein, we analyzed the behavioral response of Sitophilus oryzae to five different blends of semiochemicals found and introduced in wheat (Table 1). Briefly, these included no stimuli (negative control), UV-sanitized grain, clean grain from storage (unmanipulated, positive control), as well as grain from storage inoculated with fungal morphotype 1 (M1, identified as the asexual life stage of Aspergillus flavus) and fungal morphotype 2 (M2, identified as the sexual life stage of A. flavus). Fresh samples of semiochemicals were used for each day of testing for each assay. In order to prevent cross-contamination, 300 g of grain (tempered to 15% grain moisture) was initially sanitized using UV for 20 min. This procedure was done before inoculating grain with either morphotype 1 or 2. The 300 g of grain was kept in a sanitized mason jar (8.5 D × 17 cm H). To inoculate grain with the two different morphologies, we scraped an entire isolation from a petri dish into the 300 g of grain. Each isolation was ~1 week old and completely colonized by the given morphotype. After inoculation, each treatment was placed in an environmental chamber (136VL, Percival Instruments, Perry, IA, USA) set at constant conditions (30°C, 65% RH, and 14:10 L:D). This procedure was the same for both morphologies and was done every 2 weeks to ensure fresh treatments for each experimental assay. See file list for descriptions of each data file.

The aim of the current study was to track the movement of phosphine-resistant and -susceptible adults of the red flour beetle, Tribolium castaneum (Herbst) (Coleoptera: Tenebrionidae), which is a major pest of stored products, after brief exposures to phosphine. Exposures were followed for extended intervals to assess the recovery patterns, and how those patterns are related to known resistance to phosphine. A video-tracking procedure coupled with Ethovision software was used to assess movement after exposure. Two strains of T. castaneum were used, one susceptible and one resistant to phosphine. The susceptible T. castaneum strain had been maintained in continuous culture without any known exposure to phosphine for >30 years at the USDA-ARS Center for Grain and Animal Health Research (CGAHR), in Manhattan, KS, USA. The phosphine-resistant strain of T. castaneum was collected from wheat in Palmital, Brazil during 1988 (BRZ-5). The rearing media consisted of 95% organic, unbleached, wheat flour plus 5% brewer's yeast. Tribolium castaneum were reared under laboratory conditions of 27.5°C, and 65% relative humidity (R.H.), 14:10 L:D. Adults, of mixed sex and <1 month old, were used in the exposure bioassays. The protocol that was used in our bioassays to generate phosphine was the Phosphine Tolerance Test (Detia Degesch GmbH, Laudenbach, Germany) with some modifications, as performed by Agrafioti et al. 2021. In particular, the phosphine was generated within a plastic canister (5 L capacity) by adding 50 mL of water to two kit magnesium phosphide pellets. The concentration of phosphine gas inside the plastic canister was determined by using several dosimeter Draeger glass tubes (Draeger 25A, 0–10 000 ppm, Draeger Safety AG & Co., USA). Ten adults of each strain were placed in a plastic syringe of 100 mL with separate syringes used for each species and strain. Then, a specific gas quantity was removed from the canister with the syringe and blended with fresh air to produce a 100-mL volume with a concentration of either 1000 or 3000 ppm and compared to phosphine-free controls (0 ppm). The insects inside the syringe were held at the concentrations above for a 5 min exposure, while additional syringes containing only fresh air and insects were used as negative controls. To understand the propensity for movement after a 5 min phosphine exposure, a video-tracking procedure was used. After exposure of phosphine-resistant or phosphine-susceptible T. castaneum for 5 min, adult movement was evaluated immediately after exposure or 24 h later under the same environmental chamber conditions as the colonies (see Source Insects), but held without supplemental food. Movement was recorded for 3 h immediately after phosphine exposure but binned into 30 min intervals (e.g., 0–30, 30–60, 60–120, 120–150, and 150–180 min) in order to evaluate how movement varied over the measured time period. Movement was also recorded 24 h after exposure for periods of 1 h (binned by 30 min intervals). Movement measures of adults was tracked in six replicate Petri dishes (90 × 15 mm D:H) with a piece of filter paper (85 mm D, Grade 1, GE Healthcare, Buckinghamshire, United Kingdom) lining the bottom using a network camera (GigE, Basler AG, Ahrenburg, Germany) affixed 80 cm above the dishes. The Petri dishes were backlit using a LED light box (42 × 30 cm W:L, LPB3, Litup, Shenzhen, China) to increase contrast and affixed in place with white foam board with holes specifically cut to size for the petri dishes. Video was streamed to a nearby computer and processed in Ethovision (v. 14.0.1322, Noldus Inc., Leesburg, VA). The software was used to calculate the total distance moved (cm) and the mean instantaneous velocity (cm/s) for each adult. Each adult was considered a replicate and was never used more than once. Only adults classified as alive (normal movement speed and activity), or affected (sluggish movements or on back with legs twitching) were used in this assay. In total, 21–41 replicates were performed per treatment combination immediately after exposure, while 15–30 replicates were performed 24 h after exposure to phosphine. A total of 1525 adults were tested. There are two time periods (immediately after exposure and 24 h later), and two response variables (total distance moved in cm and mean instantaneous velocity in cm/s). There were three fixed explanatory variables: concentration of phosphine (0, 1000, or 3000 ppm), susceptibility (phosphine-susceptible or phosphine-resistant strain), and time interval (maximally 0–30, 30–60, 60–120, 120–150, and 150–180 min). Ethovision Assay morrison_ethal_ethovision_assay_fumigation_agdatacommons.csv To understand the propensity for movement after a 5 min phosphine exposure, a video-tracking procedure was used. After exposure of phosphine-resistant or phosphine-susceptible T. castaneum for 5 min, adult movement was evaluated immediately after exposure or 24 h later under the same environmental chamber conditions as the colonies (see Source Insects), but held without supplemental food. Movement was recorded for 3 h immediately after phosphine exposure, but binned into 30 min intervals (e.g., 0–30, 30–60, 60–120, 120–150, and 150–180 min) in order to evaluate how movement varied over the measured time period. Movement was also recorded 24 h after exposure for periods of 1 h (binned by 30 min intervals). Movement measures of adults was tracked in six replicate Petri dishes (90 × 15 mm D:H) with a piece of filter paper (85 mm D, Grade 1, GE Healthcare, Buckinghamshire, United Kingdom) lining the bottom using a network camera (GigE, Basler AG, Ahrenburg, Germany) affixed 80 cm above the dishes. The Petri dishes were backlit using a LED light box (42 × 30 cm W:L, LPB3, Litup, Shenzhen, China) to increase contrast and affixed in place with white foam board with holes specifically cut to size for the Petri dishes. Video was streamed to a nearby computer and processed in Ethovision (v. 14.0.1322, Noldus Inc., Leesburg, VA). The software was used to calculate the total distance moved (cm) and the mean instantaneous velocity (cm/s) for each adult. Each adult was considered a replicate and was never used more than once. Only adults classified as alive (normal movement speed and activity), or affected (sluggish movements or on back with legs twitching) were used in this assay. In total, 21–41 replicates were performed per treatment combination immediately after exposure, while 15–30 replicates were performed 24 h after exposure to phosphine. A total of 1525 adults were tested.

To determine whether colony populations of Lasioderma serricorne (cigarette beetle, CB) and Sitophilus oryzae (rice weevil, RW) vectored microbes, and to identify possible interactions with dispersal time, a vectoring assay was performed for each species. For the vectoring assay, the impact of dispersal (0, 24, or 72 h) and foraging time (3 or 5 d) on vectoring ability were tested. Briefly, adult L. serricorne or S. oryzae were singly removed from colony containers with sterilized forceps and then placed immediately in the center of Petri dish containing agar for the 0 h dispersal period. Alternatively, some insects were given a 24 or 72 h dispersal period in an autoclaved 4 L-capacity glass container and stored at constant conditions of 25°C, 60% RH, and 14:10 L:D photoperiod prior to being added to the PDA. Petri dishes were maintained at 30°C, 60% RH, and 14:10 L:D photoperiod for either 3 or 5 days, then photographed for microbial growth. Transfer of L. serricorne or S. oryzae adults from dispersal containers to agar at the conclusion of the dispersal period was performed inside the biosafety cabinet to prevent contamination of dishes. Pictures of the agar dishes and corresponding microbial growth were taken using a DSLR camera (EOS 7D Mark II, Canon, Tokyo, Japan) mounted to 3D imaging StackShot (CogniSys, Inc., Traverse City, MI, USA) equipped with a dual flash (MT-26EX-RT, Canon, Tokyo, Japan). Light was diffused using a partially cut frosted plastic jar (15.2 × 7.6 cm D:H) making a total of n = 60 replicates per treatment combination (of dispersal time, insect species, and foraging time in patch). The pictures taken were processed using ImageJ 1.53a (Wayne Rasband, National Institutes of Health, USA) to quantify the microbial growth in the agar dishes. The images had their backgrounds subtracted, then were processed using the "find edges" tool. Finally, they were converted to binary and either dilated or eroded to conform to the original image parameters. A circle encompassing the Petri dish was created and the mean grayscale, standard deviation of the grayscale value, and count of pixels was measured as a surrogate for microbial growth on the dishes. This allowed a quantitative measure of microbial growth by creating an average in a given image. The mean grayscale value could range from 0 (full white), indicating no microbial growth, to 255 (full black), indicating full microbial growth on the entire dish. Finally, visually, microbial morphospecies (alpha) richness was assigned to each image given the number of unique morphospecies on the plate as a proxy for community complexity. Treatments included those from microbially-enriched environments where Aspergillus flavus had been inoculated on wheat or flour (AF). To prepare the AF, 600 g of grain was added to a stainless-steel pot filled with water and placed on a hot plate at 500°C. Once boiling for 15 min, the water was drained and the grain was evenly spread out on sterile wipes (38.1 × 42.5 cm, 3 ply, Tech wipes, Skilcraft, NIB, Alexandria, VA) and allowed to dry inside a laminar fume hood (ca. 3 h). Afterwards, grain was evenly divided (~300 g) and placed in two separate autoclaved mason jars (950-mL capacity). A single hole was pierced through each lid and lined with a cotton ball. The jars were then sealed with aluminum foil and were autoclaved (533LS, Getinge, Rochester, NY, USA) for 30 min. To inoculate with A. flavus, a 3-inch strip of agar containing a pure culture of A. flavus grown on agar for 7 d at 30°C, 60% RH, and 14:10 L:D photoperiod was placed into each jar containing the grain. AF was then maintained at room temperature for roughly 10 d or until the A. flavus evenly covered as much the grain as possible. Batches of inoculated grain were used within 10–15 d of preparation. Grain was never used more than once for each replicate of every trial in each assay experiment to prevent cross contamination. A total of 75 insects were added to 300 g of AF in a 950-ml mason jar and allowed to forage for 2 weeks prior to use in the vectoring experiment. The same dispersal periods (0, 24, 72 h) and time in patch (3 and 5 d) described above were used for this experiment. The mean grayscale value and microbial morphospecies richness was recorded for each image. There were a total of n = 30 replicates per treatment combination. Another treatment included field-collected individuals. To obtain sufficient numbers of adults, insects were caught at four different field sites around the area of greater Manhattan, KS including: 1) a site with a pre-harvest wheat field bordered by woodlands (39°14'26.2"N, 96°34'59.1"W), 2) local apartment complex consisting of end consumers (39°11'43.6"N, 96°36'07.4"W), 3) Kansas State University Agronomy Farm with storage silos (39°12'23.7"N, 96°35'43.2"W), and 4) a private residence adjacent to a working cattle farm (39°12'23.7"N, 96°35'43.2"W). In each location, a total of three 4-funnel Lindgren traps (Bioquip, Rancho Dominguez, CA, USA) were deployed at least 10 m apart at about 1 m height on rebar or hung from a tree along the perimeter of the location site, and were baited with a multi-species lure containing both L. serricorne sex pheromone and Sitophilus spp. pheromone (PTL bullet lure, #IL-108, and Sitophilus spp. bullet lure, #IL-703, Insects Limited, Westfield, IN, USA). In addition, three ground traps were deployed that consisted of commercially-available pitfall traps (Dome®, Storgard, Trécé, Adair, OK, USA) with two connectable pieces (Doud and Phillips 2020; Doud et al. 2021), containing a central well where a Sitophilus spp. lure was added along with a 5 g of whole maize as a kairomone bait. Pheromone lures were changed every 60 d. No kill mechanism was added because adults needed to be alive. Traps were checked on a daily basis for capture of new adults and brought immediately back into the laboratory in separate unused, sterilized containers for addition to agar dishes. Stored product insects were identified using taxonomic keys in USDA (1996). Dispersal period at 0 h and time in patch (3 and 5 d) as described above were used for this experiment. Resources in this dataset: Resource Title: Full CB & RW Vectoring Dataset File Name: cb_rw_full_dataset_richness.csv

Insects Two different populations of Prostephanus truncatus were used in the bioassays, one originated from the invaded range in Ghana, and the other from the native range in Mexico. Both populations were maintained in the Laboratory of Entomology and Agricultural Zoology (LEAZ), at the Department of Agriculture, Crop Production and Rural Environment, University of Thessaly, Greece, on whole maize kernels, at 26°C and 55% relative humidity (RH) and continuous darkness. European Maize Hybrids Three different maize hybrids (“PICO”, “HAMILTON”, and “AGN 672”) were obtained from American Genetics SA, Sindos, Greece. All hybrids were cultivated at Serres, in northern Greece according to the local farming practices. The hybrid “PICO” has a great production potential and it is adapted to multiple soil types and can produce high-weight grain. The hybrid “HAMILTON” is a dual-purpose hybrid, that has excellent early vigor and it is tolerant to fungi, and the hybrid “AGN 672” has excellent early vigor and it is also tolerant to fungi. Population Growth on Different Maize Hybrids Three different maize hybrids (PICO, Hamilton, and AGN 672) were used for experimentation. These hybrids were untreated and uninfested, and kept at ambient conditions until the beginning of the experiments. Before proceeding with the bioassays, grain moisture content (M.C.) was assessed, using a moisture meter (mini-GAC plus, Dickey-John Europe S.A.S., Colombes, France). Standardized plastic vials as in prior work (Quellhorst et al. 2023; Lampiri et al. 2022) were used here (3 cm in diameter, 8 cm in height). Vials were then filled with 20 g of one of the three maize hybrids with lids added after. The commodity was weighed with a Precisa XB3200D compact balance (Alpha Analytical Instruments, Gerakas, Greece). The upper rings of the vials were treated with Fluon (Northern Products Inc., Woonsocket, USA) to prevent insects from moving away from the grain and/or escaping. The top of each vial also had small holes punched to allow ventilation. Each vial then received 10 P. truncatus adults of mixed sex and age from one of two different strains. Two different populations of P. truncatus were used as mentioned above. The vials were placed inside incubators set at 30°C and 65% R.H. in continuous darkness. The vials were removed from the incubators after 45 d and adult progeny production was recorded. We also recorded the weight of frass, the number of insect-damaged kernels (IDK), and the total weight of the kernels within each vial. For each combination, i.e. hybrid × strain, there were n = 9 replicates.

Insect Strains and Rearing Two field-derived strains of T. castaneum from either Eastern Kansas, collected in 2012, or Riley County, KS, collected in 2019, were used to assess the effect of strain on the behavioral response to necromones. Except where noted, the 2012 field strain was used for each experiment. T. castaneum was reared on a mixture of 95% unbleached flour and 5% brewer’s yeast in an environmental chamber at 27.5ºC, 60% RH, and 14:10 L:D. Subculturing proceeded by adding 75 mixed-sex T. castaneum to a 947-ml mason jar filled two-thirds with mixed diet. Adults were removed after 72 h of oviposition. Mixed sex adults aged 4–8 weeks old were used in all assays. All experiments were performed between the years 2017–2020. Treatments Time of Death of Prior Captures on Behavioral Response For investigating the attraction to kairomone oil based on how long beetles were left in the oil, the following treatments were included: negative control (neg ctrl), 950μL of Trécé Storgard® Kairomone Oil (kairomone oil for the remainder of the manuscript; Adair, OK, USA) only, or 950 μL of kairomone oil plus 25 freshly killed, mixed sex T. castaneum adults aged in the oil for 1, 25, 48, 72, or 96 h. A second round of the beetles aged longer than 8 days was included with the following treatments: negative control (neg ctrl), 950μL of kairomone oil only, or 950 μL of kairomone oil plus 25 freshly killed, mixed sex T. castaneum adults aged in the oil for 8, 9, 10, or 11 d (Table 1). These experiments were performed in a combination of the wind tunnel, release-recapture assay, and two-choice olfactometer (Table 1). Treatments were added to 20 mL GC headspace vials (Gerstel, GmBH, Germany) for wind tunnel assays, while they were added to Trécé Storgard™ Dome® traps in the release-recapture assays. Influence of Density of Prior Captures on Behavioral Response In order to evaluate whether the behavioral response of T. castaneum modulates with different densities of conspecifics in traps, the following treatments for the density response study were used: the same negative control, 950 μl of kairomone oil only, or 950 μl of kairomone oil plus either 4, 10, 20, or 40 mixed sex T. castaneum adults that were allowed to incubate for 24 h or 96 h. These experiments were performed in a combination of the wind tunnel, release-recapture assay, and headspace collection/GC-MS (Table 1). Treatments were added to 20 mL GC headspace vials (Gerstel, GmBH, Germany) for wind tunnel assays, while they were added to Trécé Storgard™ Dome® traps in the release-recapture assays. Effect of Strain on Behavioral Response to Prior Captures To rule out losing the attraction behaviors from laboratory-rearing protocols, a more recent T. castaneum strain was used and tested against the strain from Eastern Kansas collected in 2012. Thus, both a 2012 and 2019 field-collected (from Riley Co., Kansas) population of T. castaneum were tested in these experiments. The treatments for the strain effect consisted of a negative control, kairomone oil only, and 950 μl of kairomone oil plus either 4, 10, 20, or 40 mixed sex T. castaneum adults, which were allowed to incubate for 24 h. Both strains were tested in the wind tunnel and a release-recapture assay (Table 1). Effect of Rancidity on Behavioral Response to Prior Captures We conducted an experiment to test if long-term storage of the kairomone oil may have caused it to become rancid, despite being stored at 4ºC as per the manufacturer’s instructions. Treatments included: 950 μl of the kairomone oil we have used for most of our other experiments (e.g., standard Storgard® kairomone oil, or SSO) only, Storgard® kairomone oil borrowed from a colleague at the Center for Grain and Animal Health Research (CGAHR) (e.g., BSO), corn oil purchased freshly from the market (e.g., CO), or one of each of these treatments + 25 dead T. castaneum (Table 1). Attraction behavior was assessed in the wind tunnel. Assay Methods Wind Tunnel Assay Wind tunnel assays were used to evaluate upwind attraction by T. castaneum to putative necromones (e.g., see Van Winkle et al. 2022 for a description). Briefly, air was generated with a fan (diameter: 36.5 cm) connected to an inlet to the wind tunnel, where the air passed through an activated carbon filter to eliminate impurities from the air, and two successively smaller slatted-metal sieves (73 × 85 cm) to create a laminar airflow, with an average airspeed of 0.38 m/s. A purified, constant, laminar flow of air was pushed over the treatments 13.5 cm upwind of a release arena (21.6 × 27.9 cm). The odor treatments (Table 1) were positioned level with the surface of the release arena in the wind tunnel and were housed in 20 mL glass headspace vials. Caps were removed from the vials when testing commenced. The adults were placed individually in the center of the release arena and were given 2 min to make a decision, including either leaving on the stimulus edge (upwind) or a non-stimulus edge (three other edges). Adults that did not respond within the timeframe were excluded from statistical analysis. Adults were never tested more than once. All treatments were represented equally in a bout of sampling. The trials were performed inside a walk-in environmental chamber at constant conditions (27.5ºC, 60% RH), with air on purge to vent build-up of odors. Behavior was evaluated using a behavioral response index (BRI) as follows: [(T-C)/N]*100, where T is the number of adults in the treatment leaving on the stimulus edge of the arena, C is the equivalent number for the control, and N is the total sample size for both groups. The BRI can vary from 100 (full attraction) to -100 (full repellency). A total of n = 60 replicate individuals were tested, depending on assay, experiment, and treatment. Release-Recapture Assay Prior to release, 100 mixed-sex T. castaneum were settled on an 8 × 8 cm slat of cardboard for 24 h. The cardboard containing the adults was then placed in the center of a walk-in environmental chamber (5 × 6 × 2 m) set at a constant 25°C, 65% RH, and 14:10 L:D. Paper was fully laid and carefully taped on the bottom of the chamber floor to allow for easy mobility by T. castaneum. A standard Trécé Dome Trap™ that held one of each treatment (Table 1) was positioned equidistantly along the chamber’s perimeter and randomized between replicates. After 24 h, trap capture totals were calculated equal to the additional number of T. castaneum found in the trap minus those seeded in the original treatment. Experimental treatments were run simultaneously. A total of n = 8 replicates per treatment and experiment combination were used. Two-Way Olfactometer Trapping To assess preference among stimuli, T. castaneum individuals were evaluated in a two-way olfactometer. The olfactometer arena consisted of a Petri dish (9 × 1.5 cm diameter:height) with two holes drilled through opposite sides of the base at equal distances from the edge and the center of the dish. A filter paper (85 mm diameter), bisected by a faint line, was placed on the surface of the olfactometer so that the holes were on opposite sides of the filter paper (as in Morrison et al. 2020). The putative necromones (Table 1) were placed in separate, smaller Petri dishes (3.5 cm diameter) below the release arena and centered under each hole. The position of the lure and necromones was randomized between each trial. A single adult was placed in the center of the arena and left for 24 h in an environmental chamber at constant conditions (30C, 65% R.H., 14:10 L:D). A total of n = 10 replicates per comparison were performed. The percent of adults choosing each stimulus and becoming trapped in the bottom petri dish was recorded. Headspace Collection Volatiles were collected from traps seeded with 0 (oil only), 4, 20, or 40 dead T. castaneum and aged 24 h or 96 h. Central airflow was first scrubbed with a charcoal filter, then restricted to 1 L/min with flow meters. Airflow was guided through PTFE tubing to 500 mL-capacity headspace glass containers with lids and an inlet for air. The containers also had an outlet with a Porapaq-Q trap that collected volatiles for 3 h. Volatiles were then eluted with 150 µL of dichloromethane. An internal standard of 1 µL of tetradecane was also added prior to being run on the GC-MS according to standard methodology. There were n = 8 replicates per treatment. Gas Chromatography Coupled with Mass Spectrometry All headspace collection sample extracts were 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. This was coupled with a single-quadrupole Agilent 5997B mass spectrometer (MS). The compounds were separated by auto-injecting 1 μl of each sample under splitless mode into the GC-MS at room temperature (approximately 23°C). The GC program consisted of 40°C for 1 min followed by 10°C/min increases to 300°C and then held for 26.5 min. After a solvent delay of 3 min, mass ranges between 50 and 550 atomic mass units were scanned. Compounds were tentatively identified by comparison of spectral data with those from the NIST 17 library and by GC retention index. Using the ratio of the peak area for the internal standard to the peak area for the other compounds in the headspace, the emission rates of samples were normalized in ng of volatile per 950 μl aliquot of oil, per μl of solvent, and per h of collection.

Insecticide Two insecticides were used in this study: an existing formulation (tradename: Diacon IGR+ R ; Central Life Sciences, Schaumberg, IL, USA), and a new formulation with synergist (tradename: Gravista ). Diacon IGR+ contains 11.4% methoprene and 4.75% deltamethrin, with a label rate of 0.12 kg AI/L and 0.05 kg AI/L. The label rate as a residual surface treatment gives a range of 28.5 mL AI/L−171 mL AI/L H2O to cover 94 m2 for both compounds. We used the maximum labeled rate of 24 mg AI/m2 for deltamethrin and 57 mg AI/m2 for methoprene. This corresponded to 0.3 ml of the formulation in 25 ml H2O, sprayed at the rate of 0.3 ml per 50.3 cm2 arena, using an artist’s air brush (Badger 100 series, Badger Corporation, Franklin Park, IL, US) for each treatment. Each replicate was evenly applied to the concrete dish using a compressor pump. The new Gravista formulation has one labeled rate of 684 ml formulation/L H2O to cover 92.9 m2. To achieve this, we mixed 0.5 ml of the new formulation in 10 ml H2O. This was sprayed at the same rate as the other compound. Distilled water was used for the control arenas at 0.3 mL per arena. The arenas were given 8 h to dry prior to use in experiments. Insects (20 of each species per replicate) were exposed on the insecticide-treated petri dishes for either 4 or 24 h. After exposure, individual Prostephanus truncatus and Sitophilus zeamais were removed and placed into clean Petri dish arenas and evaluated for condition. Using a stereomicroscope (SMZ-18, Nikon Inc., Tokyo, Japan) under 60× magnification, P. truncatus and S. zeamais were classified as alive (moving normally, is able to right itself when flipped over, no twitching), affected (moving sluggishly or erratically, unable to right itself, twitching of antennae or legs may be present), or dead (completely immobile even after prodding) according to prior published definitions (Ranabhat et al., 2022). Dispersal and Mortality To test dispersal capacity to new food patches, a dispersal apparatus was employed. Species-specific cohorts of 20 adults (P. truncatus or S. zeamais) were exposed to Gravista, IGR+, or an untreated control as above for 4 or 24 h, then given 48 h to disperse across 30 or 70 cm standardized sections of PVC pipe (3.175 cm ID). After exposure to insecticide formulations, insects were evaluated for condition after exposure before placing them in the dispersal apparatus. The ends of both sides of the PVC pipe were sealed with mesh (425 μm) to prevent escape. At the far end of the pipe, a hole (2 cm D) was drilled and centered over a glass jar (5 × 6.5 cm D:H) to create a pitfall trap design. The glass jar contained 20 g of whole maize kernels, representing a novel food patch, to induce insects to disperse with food kairomones. Untreated, clean, and uninfested yellow maize was used in the experiments. Grain was sourced from Heartland Mills (Marienthal, KS, USA), and frozen for 72 h prior to use to ensure no prior insect infestation was present. At the end of the sampling period, the number of insects in the jar and their mortality was scored as alive, affected or dead. In addition, the position of each individual was recorded as residing in zone 1 (at the release point), zone 2 (in first half of tube), zone 3 (in second half of tube), or zone 4 (collection jar with maize). In total, there were n = 12 replicate cohorts for each species and combination of distance and treatment. In total, 1,440 P. truncatus and 1,440 S. zeamais were tested in this experiment.

Insecticide Netting In this study, we focused on two types of long-lasting insecticide netting (LLIN) that have been found to be effective for managing various stored product insect pests. One is an LLIN consisting of a polyethylene netting (2 × 2 mm mesh, D-Terrence, Vestergaard, Inc., Lausanne, Switzerland) with 0.4% deltamethrin active ingredient (a.i.), while the second one is Carifend® net (40 deniers with mesh size 97 knots/cm2; BASF AG, Ludwigshafen, Germany) containing 0.34% α-cypermethrin (a.i.). Foundational Model We used a standard Lefkovitch matrix model to project population growth for Tribolium castaneum, with four life stages (e.g., egg, larva, pupa, and adult;(Lefkovitch,1965). In equation (1), the Leftkovitch matrix L matrix (4 × 4) represents the life-stage structure of T. castaneum which has an egg, larvae, pupae, and an adult, where only the adults contribute to the fecundity, F. By multiplying L with the population vector ni(t), where t is time step (e.g., generation) and i is a life stage, we obtain the resultant vector ni(t + 1), which reveals the distribution of individuals across different life stages in the subsequent time period. In equation (1), P1 represents the probability of staying in the egg stage and G1 is the probability of moving from the egg to the larval stage, P2 is the probability of staying in the larval stage, G2 is probability of moving from the larval stage to pupal stage, P3 is the probability of staying in the pupal stage, G3 is probability of moving from the pupal stage to adult, while P4 is the probability of staying in the adult stage (Figure 1). Model Parameterization and Scenarios We simulated population outcomes for up to 15 generations by using the life table data for T. castaneum using the R package popbio. Survivorship, fecundity, and transition information for each stage were derived from the literature (summarized in Table 1). The developmental duration of eggs, larvae, and pupae were 3.82 ± 0.005, 22.81 ± 0.67, and 6.24 ± 0.071 days (Kollros,1944). The average life duration of the adult used in this study was 221.16 days (Park et al., 1961). We used 94 offspring for fertility from the study Park et al.,(1965) and 99% rate of eclosion from pupae to adult. In order to explore the sensitivity of the base model to changes in mortality and fecundity, both of these parameters were systematically varied from near zero to their maximum value given in the base model (e.g., F = 94, P4 = 0.871). The parameters were varied alone or in combination and the resulting population growth was plotted. All plots were created using ggplot2 (Wickham, 2016) in R software (R Core Team, 2022). Three empirical scenarios from the literature were modeled containing estimates of fecundity reduction only, survivorship reduction only, or both fecundity and survivorship reduction when using LLIN (R.V. Wilkins et al., 2021; Gerken et al., 2021;Scheff et al., 2021, Scheff et al., 2023; Table 2). An individual projection matrix was constructed for each of the three scenarios and combinations of the reductions in fecundity, survivorship, or both. Population growth and proportion in each life stage was projected for 15 generations for each case, including the base model. Overall variation and oscillation were calculated to compare trends among proportion of life stages in each case. In order to compare differences in population sizes between cases for all generations and for generation 15 only, population sizes for each generation were bootstrapped 1000 times to provide iterative replication. The bootstrapped data were then compared one case to another using proc ttest in SAS (Version 9.4) for all generations and for generation 15 only. In addition, a sensitivity analysis was performed to determine which stage should be targeted to most greatly affect the population growth after exposure to the netting. Moreover, a mortality function based on empirical data with LLIN exposure collected in the laboratory on T. castaneum was implemented. The three scenarios are derived from: Gerken, A. R., J. F. Campbell, S. R. Abts, F. Arthur, W. R. Morrison, and D. S. Scheff. 2021. “Long-Lasting Insecticide-Treated Netting Affects Reproductive Output and Mating Behavior in Tribolium castaneum (Coleoptera: Tenebrionidae) and Trogoderma variabile (Coleoptera: Dermestidae).” Edited by Rizana Mahroof. Journal of Economic Entomology 114 (6): 2598–2609. https://doi.org/10.1093/jee/toab204. Scheff, D. S., A. R. Gerken, W. R. Morrison, J. F. Campbell, F. H. Arthur, and K. Y. Zhu. 2021. “Assessing Repellency, Movement, and Mortality of Three Species of Stored Product Insects after Exposure to Deltamethrin-Incorporated Long-Lasting Polyethylene Netting.” Journal of Pest Science 94 (3): 885–98. https://doi.org/10.1007/s10340-020-01326-3. Wilkins, R.V., J.F. Campbell, K.Y. Zhu, L.A. Starkus, T. McKay, and W.R. Morrison. 2021. “Long-Lasting Insecticide-Incorporated Netting and Interception Traps at Pilot-Scale Warehouses and Commercial Facilities Prevents Infestation by Stored Product Beetles.” Frontiers in Sustainable Food Systems 4: https://doi.org/10.3389/fsufs.2020.561820. Resources in this dataset: Resource Title: Script for Modeling of LLIN effects on T. castaneum MS File Name: ranabhat_etal_modeling_MS_r_script_final_agdata_commons.R