Specific permission is not required for collection of BMSB annually in USDA, ARS, Beltsville Agricultural Research Center, Beltsville, MD. The land was owned by our employer, USDA. This collection will not involve endangered or protected species.

The H. halys colony was established in 2007 from adults collected in Allentown, PA, and reared on a diet of organic green beans and seeds (2:1 sunflower: buckwheat seed). Insects were maintained in Thermo Forma chambers (Thermo Fisher Scientific^®) at 25°C and 72% relative humidity with 16:8 hour light: dark cycle. The colony was replenished with ~20 field-collected bugs annually that were collected in USDA, ARS, Beltsville Agricultural Research Center, Beltsville, MD.

H. halys adults used in airborne collections were moved into separate all-male or female rearing containers two days post-imaginal ecdysis, at which point they were held until transferred to aeration containers under laboratory conditions based on methods previously developed for collecting volatiles for other insect species [14]. H. halys were placed individually (n = 116 males, n = 15 single females) or in groups (n = 89 grouped males with 6-29/group, n = 8 grouped females with 8-14/group, n = 59 third-stage nymphs with 6–20 per group in same age) in 1-liter, 4-necked glass containers with green beans and water. Volatile airborne collections were started using 5–11 day-old same sex virgin adults or 3^rd stage nymphs, and conducted in 24-hour increments for up to three months. Humidified air was drawn into the container through 6–14 mesh activated charcoal (Fisher Scientific®, Pittsburgh, PA), and out through two traps (15 cm x 1.5-cm OD) containing Super Q (200 mg each; Alltech Associates, Inc.®, Deerfield, IL) by vacuum (~1 liter/min). Insects were fed organic green beans (replaced every 2–3 days) and provided water tubes with cotton balls, and aerated continuously for 20 to 100 days depending on insect living conditions at room temperature (23–25 ⁰C) and 16L: 8D photoperiod. The adsorbent traps were changed every day (some of them in 3 days for weekend) and eluted with methylene chloride (0.5 ml/each sample). The eluents were stored in –30 ⁰C freezer until analyses.

Aeration samples were analyzed using a 30m x 0.25mm x 0.25μm inner diameter non-polar methyl silicone HP-5 capillary column (J&W Scientific Inc^®) on an Agilent 6890 gas chromatography (GC) equipped with an HP auto-sampler in splitless mode (1.4 ml/min hydrogen carrier). Oven temperature was programmed at 40°C for 2 min, then to 280°C at 15°C/min and held for 7min. A hydrocarbon, tetradecane (10 ng/μL), was used as external standard for quantitative analysis. Gas chromatography-mass spectrometry (GC-MS) analyses of semiochemicals were conducted on an HP 6890 GC equipped a HP 5973 Mass Selective Detector using same column and oven temperature programs as GC, but with helium as carrier gas (1.4 ml/min). A 70 eV electron beam was employed for sample ionization [15]. Synthetic chemical standards, tridecane, tetradecane, and E-2-decenal were purchased from Sigma Aldrich (St. Louis, MO) and Bedoukian Research, Inc. (Danbury, CT).

Male airborne collections were analyzed daily for pheromone emission as described above. When 3000 to 5000 ng/day was obtained and consistent for 2–3 days, a permeable polyethylene vial (26 mm x 8 mm x 1.5 mm thick, Just Plastic Ltd., Norwich, UK) containing 20 μL pure pheromone antagonist compound (C13 or C13+E-2-decenal in 10:1 ratio) with closed cap was placed into the aeration chamber for 24 hours. The volatile collection and analysis were continued for at least three days after the 24 hour treatment period. In a second application, 20–30 μL C13 or 20 μL C13+ E-2-decenal (10:1 ratio) was reapplied to the same chamber one week after the initial antagonist test to check for repeatability.

Individuals used in Y-tube assays were either classified as young adults (2–7 days post-adult emergence), old adults (13–23 days post-adult emergence), or 3^rd stage nymphs (13–20 days post-egg eclosion). Individuals from each group were tested in a Y-tube olfactometer for their attraction to live males. Y-tube bioassays were conducted from September-November of 2011 between 8 a.m. and 4 p.m., in a fume hood lined with black paper and covered with black curtain to prevent external visual stimuli. The Y-tube measured 30 cm for the straight tube and 15 cm for either arm, with an 8 cm diameter. At the end of each arm and the entry port for tested subjects was a modified 1L rounded flask [14]. Ambient air was pulled through an activated charcoal filter at 1 L/min and aerated with distilled water before being split through each arm. The fume hood was illuminated by a single 60 watt light bulb with a light intensity of 800 LUX, placed 50 cm above the fork in the Y-tube, with 26–28°C temperature and 50% RH. Plume structure and wind velocity were checked using incense smoke prior to the first assay.

Prior to being tested, insects were moved from Thermo Forma cages to individual 15 x 100 mm Petri dishes for one hour before being placed in the Y-tube. The Y-tube was rinsed with water and acetone after six bioassays had been conducted and if a new age group was to be evaluated. Individual insects were introduced into the release chamber and permitted to walk into either flask at the end of an arm (odor or control). If an insect did not respond within 10 minutes, it was recorded as “non-responder”. Individuals were used only once in these trials.

Attraction of nymphs, old adult males and females, and young adult males and females was observed toward live pheromone-producing males. Daily GC monitoring of samples from male aeration chambers were used to verify that males used as “Live Male” treatments in the Y-tube were actively releasing pheromone, and male lures were moved into the Y-tube 30 minutes prior to release of the test subject. A blank (empty) Y-tube was used to test H. halys movement in the assay chamber when no odors were present to ensure no bias in position.

The average and standard error of pheromone, C13, and E-2-decenal emission was calculated for single and grouped males, females, and nymphs by averaging the average/day of each aeration chamber. The start age that males began producing pheromone, C13, and E-2-decenal was calculated using average and standard error of individual males with statistics run as above. The start age by year was also calculated. Male-produced semiochemicals occurred in cyclic peaks, thus we calculated the average time in days between odor bursts, and tested whether this varied significantly by individual male using ANOVA (JMP^®).

The effect of male density on compound emission was graphed using log-transformed data and repeated-measures ANOVA was used to test the effect on each compound. The relationship between C13 and pheromone was analyzed via ANOVA pairwise comparisons. Average and standard error of compounds was calculated for single males by time of day. Time periods were standardized for number of hours collected and ANOVA was used to test for effects. Tukey HSD comparisons tested all compounds and times combined.

Y-tube bioassays were analyzed for each age group and treatment using Likelihood Ratio tests (JMP^®), with the likelihood of entering either arm set at 50% and the confidence interval set at 95%. Difference in movement towards live males was analyzed for each age/sex using Nominal Logistic Regression.

Single adult males produced more male-specific pheromone than groups of males (Table 1) while groups of males produced more C13 than single males. E-2-decenal was not a major proportion of the male odor blend. Nymphs and females did not produce pheromone. Single and grouped females and 3^rd stage nymph groups produced similar amounts of C13 and E-2-decanal. Grouped males released 10-fold more C13 than nymphs, and 100-fold more than females. Nymphs released 100-fold more E-2-decenal than males and grouped females and 10-fold more than single females (n = 116 single males; n = 89 grouped males with 6-29/group; n = 15 single females; n = 8 grouped females with 8-14/group; n = 59 third-stage nymphs with 6–20 per group).

The average and standard error of start age of the male-produced pheromone was 12.71 days ± 1.00 (n = 28). Males were placed in chambers at various ages, from 5–12 days into adult stage, but this did not affect the start age of pheromone emission. ANOVA: Age produced = Age started, year. R²-.258. F_4,27−2.00, p = 0.128. Age started (days): F-0.707, p = 0.409, Year: F-3.808, p = 0.633, n = 28. Start age (days) for pheromone in 2011: 13.29 ± 0.618 (n = 24), 2012: 11.37 ± 1.117 (n = 8); C13 2011: 13.75 ± 1.482 (n = 24), 2012: 9.00 ± 1.376 (n = 8); and E-2-decenal in 2011: 20 ± 2.540 (n = 15), E-2-decenal in 2012: 13.5 ± 3.202 (n = 4).

Cyclic patterns in emission of compounds were quantified using the numbers of days between peaks (Fig 1). Pheromone emission peaked roughly every three days. C13 and E-2-decenal did not frequently show a repeated peaking trend, rather there were one or two bursts per airborne collection. Fig 1A is an example of cyclic pheromone and random C13 peaks for a male held singly and moved to an aeration chamber five days into the adult stage. Fig 1B shows a male removed from an all-male holding cage and placed in an aeration chamber on day 9. This initial peak of C13 was frequently observed for the first 1–2 days of airborne collections if the male was not held singly. Average days between compound emission peaks are more frequent for bursts of pheromone emission relative to C13 and E-2-decenal. Average ± SE of days between pheromone peaks: 3.04 ± 0.08 days (n = 68 single males, 261 peaks); C13: 6.90 ± 0.53 days (n = 59 single males, 140 peaks); E-2-decenal: 6.64 ± 0.80 days (n = 53 single males, 140 peaks). Repeated Measures ANOVA: Square Root (Days Between Peaks) = Subject (Compound), Compound, Month: R² = 0.53; F_113,453−3.54, p <0.0001. Effect tests: Month F-0.675, p = 0.743; Compound F-44.086, p <0.0001. The average number of peaks per single male subject was: Pheromone 3.33 ± 0.27; C13 1.91 ± 0.16; E-2-decenal 0.64 ± 0.16. n = 84, Repeated Measures ANOVA: Square-root (Average number of peaks) = Subject (Compound), Compound: R² = 0.22; F_2,249−34.556, p <0.0001.

The amount of pheromone, C13, and E-2-decenal present varied by number of male H. halys adults in chamber (Fig 2). These data shows 24-hour airborne collections from 2011–2012 on 13–90 day-old adults, with log-transformed data using repeated measures ANOVA for Compound = subject, # males (subject). Pheromone F_1,71−6.634; p < 0.0001; C13 F_1,71−5.701; p = 0.001; E-2-decenal F_1,71−1.617; p = 0.003. Regression equations: Pheromone = -0.17x + 2.52; C13 = 0.22x + 1.15; E-2-decenal = -0.773x + 0.290. The relationship between C13 and the male-specific compounds was analyzed via ANOVA using pairwise correlations. Pairwise correlations, p < 0.0001. The C13 and male specific compounds interaction overlaps at 2.5 males.

Compound emission varied by time of day (Fig 3). Single males released more pheromone in the morning/early afternoon than in the evening/overnight, while C13 was released in greater amounts in the evening/overnight. Amount of compound per time period was standardized for number of hours collected. ANOVA: Log (Compound) = Time of day. For pheromone, F_1,209−35.368, p <0.0001; C13, F_1,209−5.991, p = 0.0152; E-2-decenal, F_1,209−2.919, p = 0.089 (n = 7 males, 210 time periods).

Synthetic C13 reduces pheromone emission in single males. Fig 4A is an example of C13 effect on pheromone emission for one male. The arrows indicate the date that C13 was added to aeration jars (20 μL on 10/11/12, 30 μL on 10/17/12). Fig 4B shows the average of single male pheromone emission with the application of 20 μL synthetic C13 added to aeration chambers on Day 0 (n = 10). Synthetic C13 + Aldehyde did not reduce pheromone emission in single males. Fig 5A is an example of the effect of 10:1 C13:Aldehyde (20 μL at each arrow). Fig 5B shows the average single male pheromone emission with the application of 20 μL C13 + Aldehyde on Day 0 (n = 6).

As a control, we tested the likelihood that each age group would move in an empty Y-tube (Fig 6A). For Nymphs: χ²–3.62, p = 0.05, n = 23; Young Males: χ²–0.43, p = 0.51, n = 21; Young Female: χ²–0.43, p = 0.51, n = 21; Old Male: χ²–6.06, p = 0.01, n = 21; Old Female: χ²–1.20, p = 0.27, n = 21. For the whole model, χ²–10.32, p = 0.04, DF 4, 101. Thus, nymphs are very likely to move randomly in the Y-tube and data should be interpreted with caution. Adults, especially males over 13 days into the adult life stage, are more likely to stay in the release chamber when no odor stimuli are present.

Each age group (nymphs, young adults, and old adults) was tested for attraction to pheromone-producing adult males. Twenty to 25 individuals were tested for each age. Percentage of non-responders did not differ by age group (χ²–104.38, p = 0.29, DF-4, 93); these data were not included in statistical analysis. Old adult males were the only group significantly attracted to live males in the Y-tube (Fig 6B). Nymphs: χ²–0.70, p = 0.40, n = 13; Young adult males: χ²–0.22, p = 0.63, n = 18; Young adult females: χ²–0.53, p = 0.47, n = 17; Old adult males: χ²–5.78, p = 0.02, n = 15; Old adult females: χ ²–0.08, p = 0.78, n = 13. Nominal Logistic Regression for all Ages/Sexes: χ²–3.92, p = 0.42, DF-4, 76.