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

<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 N₂ 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).

<b>昆虫</b>
本研究使用的甲虫取自美国堪萨斯州曼哈顿市的美国农业部农业研究服务局（ARS）谷物与动物健康研究中心（CGAHR）维持的种群。<i>R. dominica</i>和<i>S. oryzae</i>种群饲养于经调质至15%谷物水分的有机全麦麦粒上。传代培养时，将50只成虫置于梅森罐（容量：473 mL）内的200 mL谷物上，给予14天时间交配产卵。该时期结束后，用10号筛（2.00 mm；W.S Tyler公司，美国俄亥俄州门托市）筛除成年宿主，种群在作为寄生蜂（parasitoid）饲养宿主前需老化3周。<i>Theocolax elegans</i>分别在<i>R. dominica</i>或<i>S. oryzae</i>两种宿主上单独饲养至少三代。实验使用刚羽化、健康的<i>T. elegans</i>。所有寄生蜂种群均在独立的环境培养箱中饲养，与仅含宿主的种群分开，以防交叉污染。种群置于梅森罐中，在环境培养箱内恒定条件下培养：温度27.5℃，相对湿度60%（RH），光周期14:10（光照：黑暗）。

<b>与捕食者的相互作用</b>
实验室研究于2022-2023年在美国农业部谷物与动物健康研究中心（美国堪萨斯州曼哈顿市）开展。每年7-10月，每周从当地收获后食品设施中采集捕食者，包括堪萨斯州立大学农艺农场（GPS坐标：39.2062227，-96.5951959）——此处<i>S. oryzae</i>及其他储粮害虫大量存在（Morrison等，2025[1]）。试验中使用的大多数捕食者通过扫网（Bioquip Products公司，美国加利福尼亚州兰乔多明尼加）采集粮库附近植被，或手工采集，临时置于1加仑（=3.98 L）拉链袋中，随后用装有隔热冰袋的冷却箱立即带回实验室。实验室中，将捕食者单独放入装有10只种群来源<i>S. oryzae</i>的950 mL梅森罐中进行处理。捕食者鉴定至科水平（Marshall，2006；Paquin等，2017）。含捕食者与<i>S. oryzae</i>的梅森罐置于环境培养箱内的架子上，培养条件恒定：27.5℃，60% RH，14:10光周期。24小时后检查梅森罐，记录被取食的<i>S. oryzae</i>数量，以及<i>S. oryzae</i>是否存在聚集行为（聚集在一起并远离捕食者）——这被视为捕食者存在时非消耗性效应的证据。仅当重复次数n≥3时，捕食者的试验结果才被纳入分析。

<b>Ethovision分析</b>
采用视频跟踪结合Ethovision v.14.0软件（Noldus公司，美国弗吉尼亚州利斯堡；Noldus等，2002），研究天敌信息化合物（kairomones）对<i>R. dominica</i>和<i>S. oryzae</i>短距离移动与定向行为的影响。该系统此前已用于储粮昆虫移动与觅食行为分析（Wilkins等，2020；Ponce等，2022）。6个试验场（由培养皿构成，VWR培养皿，100×15 mm，底部用双面胶粘贴85 mm 1级滤纸，Whatman，GE Healthcare，美国芝加哥）置于网络摄像机（GigE，Basler AG，德国阿伦斯堡）下方80 cm处。每个试验场内单只昆虫的移动行为由相邻计算机同步记录。Ethovision中监测4个区域：培养皿的两个半区（处理半区vs对照半区），以及每个半区中央嵌套的两个直径1 cm的刺激区域（处理刺激区与对照刺激区）。处理位置在重复试验间随机分配，每个处理共进行n=12次重复试验。每次试验中，将单只昆虫置于试验场中央，跟踪其移动行为10分钟。Ethovision中汇总的测量指标包括：累计移动距离（cm）、瞬时速度（cm/s）、进入各刺激区的频率、进入各刺激区的潜伏期、在各刺激区的累计时间，以及在培养皿各半区的累计时间。

<b>寄生蜂信号</b>
[2]在研究寄生蜂信号对<i>R. dominica</i>和<i>S. oryzae</i>移动与觅食行为影响的试验中，采用顶空提取物（寄生蜂提取物试验）和成年寄生蜂（成年寄生蜂试验）作为刺激源。寄生蜂提取物试验的处理刺激源包括：纯净溶剂（对照）、100 g未受侵染小麦的顶空提取物（小麦组）、被<i>T. elegans</i>寄生的同种昆虫种群的顶空提取物（寄生蜂组）——各取10 μL滴加至10 mm滤纸片上。滤纸片放入试验场前，溶剂需挥发15秒。成年寄生蜂试验的处理刺激源包括：单粒小麦（小麦组）、2只雌性<i>T. elegans</i>（寄生蜂组）、无刺激源（对照组）。Ethovision试验按寄生蜂存在与否分组，以避免寄生蜂气味扩散至相邻试验场造成干扰。对照区要么无刺激源（成年寄生蜂试验），要么含滴加10 μL纯净二氯甲烷溶剂的10 mm滤纸片（寄生蜂提取物试验）。

<b>顶空挥发性物质采集</b>
为明确化学信号在介导储粮害虫与其天敌间非消耗性相互作用中的角色，采集以下样本的顶空挥发性物质：捕食者（5只蟋蟀，蟋蟀科Gryllidae）、被<i>T. elegans</i>寄生的<i>S. oryzae</i>和<i>R. dominica</i>种群、未受侵染的小麦粒。所有样本采用顶空采集系统采集（参考Van Winkle等，2022）。采用活性炭过滤器去除中央空气中的背景挥发性物质，随后气流分为8路。系统各部件通过化学惰性聚四氟乙烯（PTFE）管路和接头连接。采用在线流量计（Volatile Collection Systems公司，美国佛罗里达州盖恩斯维尔）维持每路气流速率为1 L/min。挥发性物质采集于捕集器中：捕集器为滴头式硼硅酸盐玻璃管，内含20 mg Poropak-Q吸附剂，两端由316不锈钢筛网、硼硅酸盐玻璃棉及PTFE压缩密封件固定（Volatile Collection Systems公司，美国佛罗里达州盖恩斯维尔）。采集24小时后，用氮气推动150 μL高效液相色谱级二氯甲烷（Sigma-Aldrich公司，美国密苏里州圣路易斯）洗脱捕集器中的挥发性物质。洗脱液收集于2 mL螺旋盖气相色谱（GC）瓶中（货号5191-8121，Agilent公司，美国加利福尼亚州圣克拉拉），瓶内装有带聚合物底座的150 μL玻璃内衬（货号5181-8872，Agilent公司）。所有样本用带PTFE衬里的硅胶隔垫（货号XXX，Agilent公司）磁力封盖，再用PTFE胶带密封，于-20℃保存，待用于行为试验和化学分析。

<b>化学分析</b>
取各顶空挥发性物质样本50 μL，转移至含150 μL内衬的新GC瓶中，进行气相色谱-质谱联用（GC-MS）分析。化学分析前，向每个样本中加入190.5 ng十四烷作为内标（internal standard）。样本提取物在Agilent 7890B气相色谱仪（GC）上分析，该仪器配备Agilent Durabond HP-5色谱柱（长度30 m，直径0.250 mm，膜厚0.25 μm），载气为氦气（He），流速恒定为1.2 mL/min，线速度40 cm/s；色谱仪与Agilent 5977B单四极杆质谱仪（MS）联用。分流/不分流进样口采用不分流模式，进样时温度维持在250℃。柱箱初始温度40℃，维持3分钟，随后以10℃/min升至280℃，保持3分钟。溶剂延迟5.5分钟后，扫描质量范围为35至550原子质量单位。采用C8-C20烷烃混合物计算所有峰的科瓦茨指数（Kovats index）。通过将光谱数据和科瓦茨指数与NIST 14数据库中的参考数据对比，初步鉴定各峰对应的物质。数据通过Masshunter Unknowns Analysis软件（Agilent公司，美国加利福尼亚州圣克拉拉）整理，化合物通过R包<i>uafR</i>进行比对（Stratton等，2024）。

搜集汇总

背景与挑战

背景概述

该数据集研究了寄生蜂（Theocolax elegans）和田间捕食者对两种主要储粮害虫——小谷蠹和水稻象甲的非消费性影响，重点关注行为反应和化学线索机制。实验通过视频追踪和化学分析（如顶空挥发物收集和GC-MS）方法，探究了害虫在感知天敌存在时的运动、觅食行为变化。数据集旨在为采后农业生态系统中非消费性效应的强度提供见解，数据收集于2023年至2024年在美国堪萨斯州进行。