BioControl 49: 163–185, 2004. © 2004 Kluwer Academic Publishers. Printed in the Netherlands.
A space-efficient contact toxicity bioassay for minute Hymenoptera, used to test the effects of novel and conventional insecticides on the egg parasitoids Anaphes iole and Trichogramma pretiosum Livy WILLIAMS, III∗ and Leslie D. PRICE USDA-ARS, Southern Insect Management Research Unit, P.O. Box 346, Stoneville, Mississippi 38776-0346, USA ∗ Author for correspondence; e-mail:
[email protected] Received 7 January 2002; accepted in revised form 14 July 2003
Abstract. We developed and tested a novel bioassay method for assessment of contact residues of pesticides to minute Hymenoptera. This method maintained a plant-toxin-insect interface representative of natural conditions in the field or greenhouse, and was spaceefficient. The procedure was useful in studies with both foliar residues and systemic uptake. Furthermore, the method was relatively straightforward, easy to setup, and inexpensive. Tests with the egg parasitoids Anaphes iole Girault (Hymenoptera: Mymaridae) and Trichogramma pretiosum Riley (Hymenoptera: Trichogrammatidae) confirmed that this method provided consistent, repeatable assessment of concentration-response relationships for several insecticide classes. Other researchers studying pesticide effects on minute Hymenoptera might find this bioassay method helpful, especially in situations where space is limited. We discovered differential susceptibility of these parasitoids to spinosad, thiamethoxam, and oxamyl. The order of toxicity for A. iole was spinosad>thiamethoxam>oxamyl, and for T. pretiosum was thiamethoxam>spinosad>oxamyl. Our results underscored the danger of generalizing pesticide effects across even closely related insects, and demonstrated that novel ‘selective’ insecticides are highly toxic to A. iole and T. pretiosum. Key words: Anaphes iole, biological control, Hymenoptera: Mymaridae, Hymenoptera: Trichogrammatidae, imidacloprid, parasitoids, pesticide bioassay, spinosad, thiamethoxam, Trichogramma pretiosum
Introduction Rational use of pesticides in integrated pest management relies in part on knowledge of pesticide effects on beneficial insects (Croft, 1990; Johnson and Tabashnik, 1999). Such knowledge permits strategies that minimize the disruptive effects of pesticides, such as use of selective compounds and altered rates or timings of pesticides (Way, 1986; Hassan et al., 1994;
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Martinson et al., 2001). Direct effects of pesticides, caused by direct contact with the toxin, are manifested as short-term mortality or relatively long-term sublethal effects, and generally have the greatest impact on natural enemies (Johnson and Tabashnik, 1999). Anaphes iole Girault (Hymenoptera: Mymaridae) is a minute wasp that parasitizes eggs of Lygus species (Heteroptera: Miridae) (Jackson and Graham, 1983; Udayagiri et al., 2000a). This parasitoid has potential for use in an augmentative release program for suppression of tarnished plant bug, Lygus lineolaris (Palisot de Beauvois), in cotton in the midsouth USA (Ruberson and Williams, 2000). Effective use of A. iole in an augmentative release program will depend on timing parasitoid releases so that the disruptive effects of insecticides are minimized. Because releases of A. iole will be made after insecticide applications, understanding the direct effects of insecticide residues on this wasp is critical to developing appropriate guidelines for timing releases. There is limited information about the impacts of pesticides on A. iole. Carrillo et al. (1994) reported differential susceptibility of geographically distinct populations of A. iole tested with several insecticides. Udayagiri et al. (2000b) evaluated the susceptibility of A. iole to weathered foliar residues of 14 pesticides used in California strawberry production. Several compounds compatible with inundative releases of the parasitoid were identified. We know little about the direct effects of insecticides used in cotton in the midsouth USA on A. iole. A better understanding of these impacts might permit strategies that minimize the disruptive effects of insecticides in this region. Evaluation of pesticide impacts on minute Hymenoptera is complicated by their size and extreme fragility. Previous studies have incorporated insecticides into diet (Havron et al., 1987) or have used residues on leaf or artificial substrates enclosed in cages of various types (Morse et al., 1986; Udayagiri et al., 2000b; Hassan and Abdelgader, 2001; Martinson et al., 2001; Ternes et al., 2001). Although topical application of pesticides provides important information correlating exposure to toxicity, conducting such studies with tiny insects is challenging. Ingestion of insecticide-contaminated food provides an ecologically meaningful picture of toxic effects. However, considering that many minute parasitoids frequent foliage, where they typically search for hosts, feed, mate, and rest, a bioassay evaluating the toxic effects of direct contact with residues on leaf tissue was deemed most appropriate. The IOBC/WPRS Working Group has been developing and evaluating pesticide bioassay test methods for natural enemies for more than 25 years, primarily for registration purposes and integrated control programs in
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European countries (Sterk et al., 1999; Candolfi et al., 2000). A standardized hierarchical testing scheme (laboratory, semi-field, and field tests) provides the necessary information for safe and effective use of pesticides and permits the development of internationally approved guidelines. It is recognized that no single bioassay method provides definitive and complete information to assess the impacts of pesticides on insects (Hassan, 1989; Rowland et al., 1991), and that development and use of test procedures is driven in part by logistical realities as well as by the specific objectives of the research. Our objective was to develop a bioassay method by which we could assess foliar residues of insecticides on acute mortality of A. iole. This would facilitate studies of field-weathered residues that might lead to refinement of timing of parasitoid releases so that pesticide-induced mortality is reduced. A logistical constraint was that the bioassay method be space-efficient. It was also important that the method be versatile, i.e., that it could be used with different means of pesticide application (leaf dip, sprays, and systemic), and with other species of minute parasitoids. Other important considerations included consistent results, relative ease of set up, and minimal expense. The new bioassay method was used to assess the toxicity of novel and conventional insecticides to A. iole and Trichogramma pretiosum Riley (Hymenoptera: Trichogrammatidae), a widespread and important egg parasitoid of several lepidopterous pests (Hoffmann et al., 1990; Li, 1994).
Materials and methods Insects Anaphes iole used in this study were obtained from a laboratory colony at the USDA-ARS BCMRRU, Starkville, MS, and were maintained on host (Lygus hesperus Knight) eggs. Trichogramma pretiosum were purchased from Buglogical, Inc., Tucson, AZ, and were reared on Sitotroga cerealella (Oliver) (Lepidoptera: Gelechiidae) eggs. Both suppliers provided parasitized host eggs which were held in Plexiglass cages (26 cm × 26 cm × 20 cm) at 27 ± 1 ◦ C, 65–85% RH, and 14:10 L:D photoperiod. Upon emergence, wasps were provided ad lib with distilled water and a 1:1 mixture of honey:distilled water for 3 days (A. iole) or 1 day (T. pretiosum), at which time bioassays were conducted. Bioassay chamber Each bioassay chamber consisted of the following components. One piece of flexible transparent tubing (VWR cat. no. 60985-584) (2.54 cm ID, 3.5 cm
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(a)
(b) Figure 1. (a) Components of a bioassay chamber: (1) one piece flexible transparent PVC tubing (2.54 ID × 3.5 cm long) with four ventilation holes (1 cm diam), (2) two pieces nylon organdy (3.5 cm × 2 cm), (3) two scintillation vial caps each filled with 1.5 ml agar gel, (4) one piece dialysis membrane (12 cm × 2 cm), (5) one tip (1.5 cm long) cut from plastic pipet tip. (b) An assembled bioassay chamber.
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long). A cork borer was used to cut four ventilation holes (1 cm diam) in the center of the tube at equidistant spacing. Two of the holes on opposite sides of the tubing were covered with pieces of nylon organdy (3.5 cm × 2 cm) which were affixed to the outside of the tube with a small amount (ca. 0.8 cm × 0.8 cm) of water-soluble, non-toxic glue (Elmer’s Washable School Glue Stick, Elmer’s Products, Inc.). Glueing the organdy to the outside of the tube facilitated subsequent assembly of the bioassay units. Two caps for scintillation vials (Kimble Glass Inc., cat. no. 74521-22400) each filled with ca. 1.5 ml agar gel (1.5% by weight). One piece (12 cm × 2 cm) of dialysis membrane (Daigger Inc., cat. no. VISMC-1-778X500). One tip 1.5 cm long cut from a Finntip 5–300 µl pipet tip (Labsystems, Inc., cat. no. 9401250). The components of the bioassay chamber are shown in Figure 1a. Bioassay procedure Bioassay chambers were set up in the following manner. Cotton, Gossypium hirsutum L. (var. DPL NuCOTN 33B), seedlings were grown in plastic flats in an insect-free greenhouse. At the 2 true leaf stage leaf disks (2.3 cm diam) were excised from the true leaves. The leaf disks were dipped for 20 s in either formulated insecticide diluted with distilled water (treatments) or distilled water only (controls). Solutions were prepared immediately before use. The leaf disks were allowed to air-dry, after which they were placed individually on the agar gel in each scintillation vial cap. Prior to this, approximately 20 µl of chicken egg albumin was placed on the gel in each vial cap. One leaf disk was placed with the upper surface exposed, and the other disk with the under surface exposed. Agar prevented the leaf disks from wilting during the bioassay, and the egg albumin acted as an adhesive, ensuring that the leaf disks remained tightly sealed to the agar. The chambers were assembled by sliding one cap into each end of the transparent tubing so that the edge of the leaf disk was aligned with the edges of the ventilation holes. The under surface of the leaf disk formed the chamber’s ceiling, while the upper surface of the other leaf disk formed the floor, thus approximating natural conditions in the field. A moistened strip of dialysis membrane was then wrapped around the ventilation holes at the center of the chamber and allowed to air dry. Upon drying, the membrane shrunk so that the caps with leaf disks were held firmly in position. A pointed scalpel (Feather Safety Razor Co., Ltd., no. 11) was then used to cut a circular piece of dialysis membrane away from the organdy-covered ventilation holes. A piece of borosilicate glass capillary (World Precision Instruments, Inc., cat. no. 1B150F-4) ca. 5 cm long, 1.5 mm diam, and flame-drawn to a point at one end was then used to make a hole in one of the membrane-covered ventilation holes. Through this hole parasitoids (mixed gender, assumed mated) were introduced to each bioassay chamber. The number of wasps introduced
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to each chamber varied from ca. 20 to 25 depending on the trial. This was accomplished using an aspirator constructed from a 2-cm long piece of glass capillary of the same type used to make the hole. The 1.5 cm long piece of pipet tip was then inserted snugly into the hole and ca. 25 µl of a 1:1 honey:distilled water mixture was added. An assembled bioassay chamber is presented in Figure 1b. Bioassay chambers were then placed in polypropylene racks (VWR, cat. no. 60982-060) arranged by replicate and held in an environmental chamber at 27 ± 1 ◦ C (A. iole) or 22 ± 1 ◦ C (T. pretiosum), 65–85% RH, and 14:10 L:D photoperiod. The air in the environmental chamber was circulated continuously by two 15-cm diameter fans. After the exposure period (A. iole, 48 h; T. pretiosum, 24 h), mortality was assessed by observing wasps in each chamber with a dissecting scope (10–50×). The exposure period for T. pretiosum was limited to 24 h because longevity of this species is ca. three days when provided with a honey diet (Ruberson and Kring, 1993). Wasps not exhibiting repetitive (non-reflex) movement were scored as ‘dead’. Our observations indicated that ≤5% of the wasps were trapped in the feeding tubes; these wasps were not included in the data analysis. Following mortality assessment, bioassay chambers were disassembled and the transparent tubing, nylon organdy, and pipet tip were washed in hot water and Alconox , triple-rinsed in distilled water, and air-dried. The remaining components were discarded. Monetary costs, setup time, and other resource constraints associated with this bioassay method were recorded. Direct contact versus fumigation Initial studies were conducted with A. iole to verify that the bioassay method assessed direct contact toxicity, not a combination of fumigation and contact toxicity. In the first study we chose seven insecticides based on their widespread use in agriculture and diverse chemical classification. This study was conducted with field rates assuming a coverage rate of 93.5 l/ha. The insecticides tested were oxamyl [Vydate 2 emulsifiable concentrate (EC), carbamate, (0.28 kg ai/ha); E. I. DuPont de Nemours & Co., Wilmington, DE], spinosad [Tracer 4 suspension concentrate (SC), bacterial fermentation, (0.1 kg ai/ha); Dow AgroSciences, Indianapolis, IN], cyfluthrin [Baythroid 2 EC, pyrethroid, (0.045 kg ai/ha); Bayer, Inc., Kansas City, MO], lambda-cyhalothrin [Karate 1 EC, pyrethroid, (0.034 kg ai/ha); Syngenta, Wilmington, DE], thiamethoxam [Actara 25 wettable granular (WG), neonicotinoid, (0.1 kg ai/ha); Novartis, Greensboro, NC], fipronil [Regent 2.5 EC, phenyl pyrazole, (0.043 kg ai/ha); Aventis, Research Triangle Park, NC], and acephate [Orthene 75 soluble powder (SP), organophosphate, (0.28 kg ai/ha); Valent USA Corp., Walnut Creek, CA].
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Insufficient numbers of wasps precluded a single trial with all insecticides; therefore a separate trial was conducted for each insecticide. Approximately 20 wasps were used in each bioassay chamber. Each trial was setup in a completely randomized design with the following four treatments. For one treatment, chambers were assembled as described above, with leaf disks exposed. Mortality in this treatment could be due to toxicity via direct contact as well as fumigation effects, if any. Another treatment was setup with a piece of nylon organdy (6 cm × 6 cm) placed over both leaf disks prior to insertion of the vial caps into the tubing. This organdy barrier did not allow wasps direct contact with the insecticide residues on the leaf disks; thus mortality in this treatment could be attributed solely to fumigation. The remaining two treatments were controls (leaf disks dipped in distilled water) assembled in the same way (with or without organdy barriers covering the leaf disks) as the first two treatments. Seven to 10 replicates were used for each treatment. Another study was conducted to evaluate contact vs. fumigation effects in greater detail. In particular, our objective was to evaluate the importance of ventilation of the bioassay chambers on possible fumigation effects, and thereby determine if a simpler window (dialysis membrane) might be acceptable for use with the bioassay chambers. This study was setup in a completely randomized design with the following eight treatments. Four control treatments were setup, two as described above, i.e., with or without organdy barriers over the leaf disk and both treatments with organdy windows for ventilation. The remaining two control treatments were also setup with or without organdy barriers over the leaf disk but all of the windows were covered with dialysis membrane. The dialysis membrane restricts ventilation more than organdy, but does allow enough air circulation to avoid condensation that would trap the wasps. Using only dialysis membrane for the windows would permit more rapid assembly of the chambers. For the four insecticide treatments the bioassay chambers were assembled in the same manner, i.e., with/without organdy barriers and both types of ventilation. Treatments were replicated four to eight times. We used acephate and imidacloprid [Provado 1.6 flowable (F), chloronicotinyl, Bayer, Inc., Kansas City, MO], at 100 and 1000 µg ai/ml, respectively. These concentrations are ca. 0.033× (acephate) and 1.06× (imidacloprid) the field rates, and in preliminary studies resulted in >85% mortality. Approximately 24 wasps were used in each bioassay chamber. The factorial treatment design allowed us to partition the importance of vapor effects and ventilation of the bioassay chambers on wasp mortality. Systemic uptake A study was conducted to determine if the bioassay chamber was useful for evaluation of systemic insecticides against natural enemies. Cotton seedlings
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were cut at the mainstem near the soil line and placed immediately in the desired concentrations of imidacloprid (32 µg ai/ml) [Admire 2 flowable (F), chloronicotinyl, Bayer, Inc., Kansas City, MO], or thiamethoxam (16 µg ai/ml). Seedlings chosen as controls were placed in distilled water. All the seedlings were then placed in an environmental chamber (27 ± 1 ◦ C, 65– 85% RH, and 24:00 L:D photoperiod) for 24 h hydroponic uptake. Leaf disks were then cut and bioassay chambers were assembled (leaf disks exposed, with organdy windows) as described in the ‘bioassay chamber’ section. At this time, untreated leaf disks dipped in the same concentrations of these insecticides (or distilled water) were also setup, thus facilitating comparison of mortality via systemic uptake vs. leaf dip. Approximately 19 A. iole were used in each bioassay chamber. Eight bioassay replicates were setup for each treatment. Concentration-response After verification that the bioassay method assessed direct contact toxicity, a study was conducted to estimate the toxicity of oxamyl, spinosad, and thiamethoxam to A. iole and T. pretiosum wasps. Bioassay chambers were assembled with leaf disks exposed and with organdy windows. Dilution series were made using formulated insecticide and distilled water on the basis of µg active ingredient per ml of solution. For each insecticide, five concentrations were chosen based on preliminary studies that established the range of mortality for these compounds. Spinosad was tested at concentrations ranging from 0.1 to 1.0 µg/ml for both species. Thiamethoxam was tested at concentrations ranging from 0.1 to 3.2 µg/ml for T. pretiosum, and 1.0 to 32 µg/ml for A. iole. Oxamyl was tested at concentrations ranging from 3.0 to 210 µg/ml for T. pretiosum, and 32 to 320 µg/ml for A. iole. A control treatment consisted of leaf disks dipped in distilled water. Approximately 23 wasps were introduced into each chamber. Separate trials were setup for each species-insecticide combination, and trials were repeated two to six times for these combinations. Each trial was setup using four to eight bioassay chambers (subsamples) per treatment. Data analysis Direct contact versus fumigation For the study with field rates of seven insecticides, low numbers of male wasps precluded their analysis; thus only data for female wasps were analyzed. Data for wasp mortality were arcsine-square root transformed using an adjustment for small proportions (formula 13.8 in Zar, 1996) prior to single factor analysis of variance by PROC GLM (SAS Institute, 2000). Extreme
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mortality (100%) in the insecticide without organdy treatment led to a lack of homogeneity of variance; therefore this treatment was omitted from analysis of variance. Analysis of variance F-tests and least significant difference tests were used to compare controls with organdy barriers vs. insecticides with organdy barriers and controls with organdy barriers vs. controls without organdy barriers (α = 0.05). Retrospective power tests (α = 0.05) were conducted to assess the precision of the study, i.e., to determine the likelihood of obtaining a statistically significant result given that a biologically significant difference in fact existed. For the study evaluating ventilation of the bioassay chambers and vapor effects of acephate and imidicloprid, wasp mortality was arcsine-square root transformed as described above prior to 4-way crossed factorial analysis of variance by PROC GLM (SAS Institute, 2000). Sufficient numbers of female and male wasps allowed the inclusion of gender in the model. High mortality (>80%) in the insecticide treatments without organdy barriers over the leaf disks did not cause a lack of homogeneity of variance; therefore these treatments were included in the analysis of variance. Analysis of variance F-tests and least significant difference tests were used to compare treatment means (α = 0.05), and power tests (α = 0.05) were conducted as described above. Mortality data from the systemic uptake study were arcsine-square root transformed as described above prior to 3 × 2 factorial analysis of variance by PROC GLM (SAS Institute, 2000). Only female A. iole were present in sufficient numbers for analysis. Analysis of variance F-tests and least significant difference tests were used to compare treatment means (α = 0.05). For the concentration-response study, low numbers of male wasps precluded their analysis; thus only data for female wasps were analyzed. Concentration-mortality data were control-adjusted and analyzed using probit analysis, including tests of parallelism (equal slopes) and equality (equal slopes and intercepts) (LeOra Software, 1987). Comparison of lethal concentrations (Robertson and Preisler, 1992) were then calculated: (1) between species for each insecticide, and (2) between insecticides for each species. Significant differences were indicated when the 95% CI of a lethal concentration ratio did not bracket the value 1.0.
Results and discussion Direct contact versus fumigation Results of single factor analysis of variance excluding the insecticide without organdy treatment demonstrated that mortality of A. iole did not differ (P > 0.05) for the seven insecticides tested at field rates (Table 1). Mortality was
0.043 0.28 0.28 0.1 0.1 0.045 0.034
Fipronil Acephate Oxamyl Spinosad Thiamethoxam Cyfluthrin L-cyhalothrin
305 336 590 269 502 270 246
No. waspsb
5.59 5.51 13.1 17.2 13.9 17.2 8.45
7.30 14.6 20.1 27.4 16.2 14.9 19.2
6.11 6.08 16.8 21.3 14.0 22.5 21.2
100 100 100 100 100 100 100
Treatment mortality (mean%)a With organdy Without organdy Control Insecticide Control Insecticide 16 18 22 22 20 23 20
Error dfc
0.31 1.36 1.22 0.77 0.36 0.51 1.88
Fc
0.7364 0.2807 0.3139 0.4743 0.7018 0.6074 0.1783
Pc
0.813 0.638 0.900 0.448 0.951 0.446 0.540
Power (1 − β)d
6.71 8.17 5.57 10.7 4.90 10.8 9.34
LSDe
‘control with organdy’ vs. ‘control without organdy’ treatments were not significant (P > 0.05).
a Untransformed data. b Number of female wasps used in analysis of variance. c Error df, F, and P values for single factor ANOVAs excluding ‘insecticide without organdy’ treatment due to lack of homogeneity of variance. d The power, using the harmonic mean, of detecting a difference of ≥10% if it exists, α = 0.05. e Least significant difference for one-tailed test, α = 0.05. All comparisons between ‘control with organdy’ vs. ‘insecticide with organdy’ and between
Rate (kg ai/ha)
Insecticide
Table 1. Mortality and statistics for contrasts testing for fumigation effects of insecticides on female Anaphes iole Girault
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Table 2. Mortality of Anaphes iole Girault wasps in bioassay chambers with variable ventilation and exposure to acephate and imidacloprid Organdy Type of barrier window
With With Without Without
Treatment mortality (mean%)a SEa No. Control Acephate Imidacloprid waspsb (distilled water) (100 µg ai/ml) (1000 µg ai/ml) Female Male Female Male Female Male
Organdy 7.55 Dialysis 7.99 Organdy 8.75 Dialysis 12.3
5.56 17.6 14.5 14.0
6.67 10.9 98.7 97.4
0.00 3.85 1.92 14.5 12.5 23.6 81.3 97.2 100 89.5 100 100
6.64 4.69 6.64 4.69
229/77 434/172 222/72 374/141
a Untransformed data ((least squares means and standard errors adjusted for other terms in
the model (SAS Institute, 2000)). b Number of wasps (female/male) that were used in analysis of variance.
high (>98%) when wasps were subjected to insecticides without organdy barriers. High mortality in this treatment confirmed that insecticide rates were high enough to provide an adequate test of contact vs. fumigation effects. Comparisons between treatments indicated that mortality of female A. iole in insecticide treatments with organdy barriers was not significantly greater than in controls with organdy barriers (P > 0.05) (Table 1). In fact, cyfluthrin caused numerically less mortality than was observed in controls. Mortality between control treatments (with vs. without organdy barriers) did not differ significantly (P > 0.05), indicating that the nylon organdy did not contribute to mortality of the wasps. Mortality in controls with organdy barriers ranged from 5.51 to 17.2%, while mortality in insecticide treatments with organdy barriers ranged from 7.30 to 27.4%. Power values for trials with fipronil, acephate, oxamyl, and thiamethoxam were sufficient to indicate that these trials had adequate precision. However, we observed higher variability in the trials with spinosad, cyfluthrin, and L-cyhalothrin, which in turn, led to lower power and the increased possibility of real differences of ≥10%. Results of factorial analysis of variance revealed that different treatments influenced mortality of A. iole in the study with acephate and imidacloprid (F = 48.49; df = 23,118; P < 0.0001) (Table 2). Mortality was high (>80%) when wasps were subjected to either insecticide without organdy barriers, regardless of the type of ventilation window. Mortality of female and male A. iole in insecticide treatments with organdy barriers was not significantly greater than in controls with organdy barriers (P > 0.05) regardless of the type of ventilation (Table 2). In fact, mortality of female and male wasps in the organdy barrier with organdy window treatment was numerically less for both insecticides than in the corresponding control treatment.
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Figure 2. Effect of insecticide treatment and differential exposure (direct contact vs. putative fumigation) on mortality of A. iole.
Figure 3. Effect of gender and differential exposure (direct contact vs. putative fumigation) on mortality of A. iole.
We observed a significant insecticide by barrier interaction (F = 117.72; df = 2,118; P < 0.0001) (Figure 2). The power test for this interaction gave a value of 0.999. In the insecticide treatments wasp mortality was significantly reduced (P < 0.0001) when the organdy barrier was present, but the presence of the barrier did not affect mortality in the control treatment (P = 0.2973). In a comparison of insecticide treatments with organdy barriers, no signifi-
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cant differences in mortality were observed between acephate and the control (P = 0.9872) or between imidacloprid and the control (P = 0.9041). These results indicate that contact toxicity is the most important factor leading to mortality of A. iole wasps exposed to acephate and imidacloprid. We also observed a significant barrier by gender interaction (F = 5.16; df = 1,118; P < 0.0250) (Figure 3). The power test for this interaction gave a value of 0.615. In the absence of organdy barriers female wasps suffered numerically greater mortality than did males (P = 0.0697). When barriers were present, mortality of male wasps was numerically greater than for females (P = 0.1705). For both genders mortality was greater (P < 0.0001) in the absence of barriers than in their presence. The type of ventilation window also had a significant effect on wasp mortality (F = 7.99; df = 1,118; P < 0.0055). The power test for this main effect gave a value of 0.801. Comparison between type of window across treatment and gender showed that percent mortality (mean ± SE) of A. iole was greater in bioassay chambers with dialysis membrane windows (41.7 ± 1.37) than in chambers with organdy windows (35.5 ± 1.92). It is possible that the relatively still air inside the bioassay chambers with dialysis membrane windows was not conducive to survival of A. iole. For this reason, in subsequent trials we used bioassay chambers of the original design (with two organdy windows), although with proper validation it may be acceptable to use windows entirely of dialysis membrane in investigations with other natural enemies. Results from the two studies assessing contact vs. fumigation indicated that fumigation effect was not a significant mortality factor for A. iole in the bioassay chambers, and thus mortality could be attributed to direct contact to the selected insecticides. That these results were consistent for seven different classes of insecticides suggests that the bioassay chamber may be useful for evaluating contact toxicity of a broad range of insecticides. Of course, to be prudent future studies with other compounds should include trials evaluating fumigation vs. contact toxicity. When fumigation effects are found to be present, ventilation of the bioassay chambers can be increased by adding more ventilation windows to the chambers, or by the use of additional fans or pumps (Morse et al., 1986; Haseeb et al., 2000). However, in cases where fumigation effects are not an issue, such as in the present study, avoiding these measures conserves resources (e.g., money, laboratory space, and setup time). These trials represented a ‘worst-case’ scenario for fumigation effects, i.e., high insecticide rates and no delay between application and exposure to natural enemies. Therefore, it stands to reason that in investigations using lower concentrations and/or longer intervals between application and exposure, fumigation effects of these insecticides will not be a concern. Our results confirmed that this bioassay method provides an
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Figure 4. Effect of imidacloprid and thiamethoxam on mortality of female A. iole (n = 812) when applied as hydroponic uptake (systemic toxicity) or leaf dip (direct residual contact).
appropriate assessment of contact toxicity to insecticides comprising a wide range of chemical classes. Systemic uptake Results of factorial analysis of variance indicated that mortality of A. iole wasps was significantly affected by exposure to cotton leaves treated by either systemic uptake or leaf dip with imidacloprid and thiamethoxam (F = 90.77; df = 5,42; P < 0.0001). Significant differences in percent mortality (mean ± SE) were observed between imidacloprid (86.5 ± 2.01), thiamethoxam (92.9 ± 2.01), and controls (11.4 ± 2.01) (F = 224.8; df = 2,42; P < 0.0001; LSD = 5.62). Mortality for each treatment-application method combination is presented in Figure 4. These results indicate that this bioassay method is suitable for investigations of insecticides with systemic properties as well as non-systemic compounds. It is especially interesting to note the high toxicity to A. iole resulting from systemic application of these insecticides, because systemic insecticides are generally less toxic to beneficial insects than other compounds. Systemic application of oxamyl, aldicarb, and disulfoton did not adversely affect parasitism of Liriomyza trifolii (Burgess) by the braconid Oenongastra microrhopalae (Ashmead) (Oetting, 1985). Soil application of dimefox or mephosfolan controlled early-season populations of Phorodon humuli (Schrank), without affecting subsequent predation by Anthocoris nemoralis (L.) (Solomon, 1987). Although systemic insecticides
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offer great potential for pest control while minimizing deleterious effects on beneficial insects, it is clear that such is not always the case; therefore detailed studies must be done on a case-by-case basis. Concentration-response Probit analysis estimated LC50 and LC90 values for female A. iole and T. pretiosum wasps subjected to contact residues of oxamyl, spinosad, and thiamethoxam (Table 3). All comparisons of lethal concentrations between insecticides for each species were significant (P < 0.05). For A. iole, the order of toxicity was spinosad>thiamethoxam>oxamyl. This relationship was consistent between LC50 and LC90 estimates. For T. pretiosum, the order of toxicity for the LC50 estimate was spinosad>oxamyl, and for the LC90 was thiamethoxam>spinosad>oxamyl. Comparisons of lethal concentrations between species for each insecticide must be done with care because exposure times were different for Anaphes (48 h) and Trichogramma (24 h). However, the trends in toxicity suggest significant differences between the species for spinosad and thiamethoxam treatments. For spinosad, the LC50 value for A. iole (0.50 µg/ml) was greater than for T. pretiosum (0.23 µg/ml). This difference probably would have been even greater if the exposure time for A. iole had been 24 h, or conversely, if the exposure time for T. pretiosum had been 48 h. The LC90 estimates were not significantly different between the two species. This was due to converging slopes at higher concentrations, which indicates a change in toxicity per unit change in concentration. For thiamethoxam, the LC90 value was greater for A. iole (17.6 µg/ml) than for T. pretiosum (0.53 µg/ml). Again, this difference would probably have been greater if exposure times had been equivalent. All slope and line comparisons between insecticides for each species were significant. These results suggest that the manner(s) of detoxification (e.g., target site sensitivity, absorption through the cuticle, and excretion) differ between these compounds for A. iole and T. pretiosum. Comparisons between species for the same insecticide indicated slopes and lines were different for spinosad and thiamethoxam, but not for oxamyl. This suggests that A. iole and T. pretiosum detoxify oxamyl in a similar manner, but that detoxification of spinosad and thiamethoxam differs between these species. Toxic effects of a given insecticide, including novel compounds purported to be ‘softer’ on beneficial insects than conventional insecticides, can differ greatly between parasitoid species. For example, our results indicated that A. iole and T. pretiosum differed in their susceptibility to residues of the two novel compounds tested, spinosad and thiamethoxam. Moreover, both compounds were more toxic to these parasitoids than oxamyl. Conversely, Elzen et al. (1999) found spinosad to be significantly less
Oxamyl Spinosad Thiamethoxam Oxamyl Spinosad Thiamethoxam
A. iole A. iole A. iole T. pretiosum T. pretiosum T. pretiosum
58.60 (34.2–85.2) 0.50 (0.38–0.58)∗ 1.70 (0.83–2.75) 52.70 (32.2–76.6) 0.23 (0.15–0.30)∗ Nd
LC50 µg/ml (95% CI)a 229.00 (149–523) 1.03 (0.87–1.41) 17.60 (9.56–55.71)∗ 223.00 (166–354) 1.19 (0.85–2.07) 0.53 (0.39–0.77)∗
LC90 µg/ml (95% CI)a
No. waspsc 1818 2725 3413 924 3410 3447
Slope ± SEb 2.16 ± 0.13 4.01 ± 0.29† 1.27 ± 0.05† 2.15 ± 0.16 1.78 ± 0.09† 0.85 ± 0.07†
130 185 471 20.1 236 47.6
χ2
13 28 26 5 25 20
df
confidence limits for each ratio (Robertson and Preisler, 1992)). All LC comparisons between insecticides for each species are significant. b Values with ‘† ’ denote slopes that are significantly different between species for each insecticide (determined by tests of parallelism (Robertson and Preisler, 1992)). All slope comparisons between insecticides for each species are significant. c Number of female wasps used in regression analysis. Nd = not determined.
a Values with ‘∗ ’ denote LC estimates that are significantly different between species for each insecticide (determined by estimation of 95%
Insecticide
Species
Table 3. Toxicity of selected insecticides to female Anaphes iole Girault and Trichogramma pretiosum Riley wasps
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toxic to Catolaccus grandis than oxamyl. Tillman and Mulrooney (2000) also reported that toxicity of spinosad residues varied widely between the braconids Bracon mellitor Say, Cardiochiles nigriceps Viereck, and Cotesia marginiventris (Cresson), although spinosad was generally less toxic than lambda-cyhalothrin. Pietrantonio and Benedict (1997) suggested that spinosad residues were more toxic to Cotesia marginiventris than to C. plutella. Collectively, these findings illustrate that even novel insecticides are differentially toxic to closely related parasitoid species. This underscores the importance of determination of pesticide effects on particular species, and not generalizing across taxa, guilds, or other groups. As is the case with many natural enemies, A. iole and T. pretiosum are more susceptible to certain insecticides than to others. Our results extend the knowledge of differential insecticide impacts on these parasitoids. Carrillo et al. (1994) reported that A. iole wasps from different geographic regions of the western United States exhibited differential susceptibility to methamidophos, endosulfan, and formetanate. Susceptibility of A. iole to oxamyl, spinosad, and thiamethoxam in the present study was intermediate between formetanate, and methamidophos and endosulfan in the study of Carrillo et al. (1994). Udayagiri et al. (2000b) evaluated the effect of residue age of 14 pesticides used in California strawberry production on toxicity to A. iole. Of the six insecticides tested, foliar residues of fenpropathrin, bifenthrin, and carbaryl were more toxic (>75% mortality 13 days after treatment) than residues of malathion, methomyl, or naled (50% mortality ≤13 days after treatment). Stinner et al. (1974) reported ca. 75% mortality of T. pretiosum up to 1.6 km downwind from ULV applications of methyl parathion. Greenhouse-weathered residues of permethrin were significantly more toxic to T. pretiosum than residues of endosulfan (Jacobs et al., 1984). In a review of the literature, Bull and Coleman (1985) reported differential insecticide toxicity across Trichogramma species, and presented data demonstrating that residues of methyl parathion and methomyl were significantly more toxic to T. pretiosum than were residues of dimethoate, toxaphene, chlordimeform, phosmet, and permethrin. Kring and Smith (1995) reported that foliar residues of thiodicarb and lambda-cyhalothrin were both highly toxic to T. pretiosum for at least 5 days after application. The present study documents the first reports of acute mortality of A. iole and T. pretiosum for several novel insecticides. More detailed assessments of these and other novel insecticides are warranted, especially evaluations of sublethal and indirect effects (Johnson and Tabashnik, 1999). We developed a novel bioassay method to facilitate the evaluation of pesticide effects on minute Hymenoptera. The method satisfies our requirements for a procedure that assesses direct contact toxicity; thus it may prove useful
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to other researchers. The method has several practical attributes, including being relatively straightforward and easy to setup. Importantly, the method is space-efficient; 100 bioassay chambers require only 0.5 m2 (ca. one shelf in an environmental chamber). The method also permits easy recovery of organisms that survive tests so that sublethal and/or indirect effects can be evaluated. Resource constraints are also relatively low. For example, initial supply costs for 100 bioassay chambers is ca. US $100. Of this cost, the reusable and discarded supplies have costs of US $60 and $40, respectively. Time required to setup 100 bioassay chambers is ≤16 person/hours. Mortality assessment, disassembly, and washing the chambers require ≤8 person/hours. We used plant tissue (leaf disks) in this method because doing so maintains the plant-toxin-insect interface and thus more closely represents pesticide exposure to parasitoids in the field. The use of leaf tissue also enhances the versatility of the procedure, making possible more detailed work while bridging the gap between laboratory and field studies. Bioassay methods that do not incorporate plant tissue generally do not permit such work. Our bioassay method provided well-defined concentration-response relationships for different insecticide classes; due in part to the fact that the bioassay chambers enclosed a high proportion of treated area, thus making exposure more consistent. Although we used leaf disks dipped in insecticide solutions, the bioassay method is also suitable for use with foliar residues applied by Potter Precision Laboratory spray tower (Burkhard Manufacturing Co., Ltd., Hertfordshire, UK), by hand sprayers, or by commercial field sprayers. In addition to facilitating studies of foliar residual toxicity, the bioassay method also has application for studies of systemic compounds. In fact, the bioassay method may prove useful in separating residual vs. systemic effects of insecticides with both properties, e.g., imidacloprid and thiamethoxam. Application of pesticides by leaf dip is widely accepted in studies with beneficial and pest arthropods (Rowland et al., 1991; Bellows and Morse, 1993; Cahill et al., 1996a; Martinson et al., 2001). Advantages of this technique include ease of setup and minimal requirements for equipment. A disadvantage is that leaf dip residues may be uneven, leading to increased variability in response (Dittrich, 1962). As an alternative to leaf dips, the Potter spray tower (Kolmes et al., 1991; Jansen, 2000; Tillman et al., 2001) applies a specified amount of insecticide per unit area, but requires specialized and expensive equipment to which many laboratories do not have access. A third approach, systemic application, is accomplished by either hydroponic uptake or soil application, and facilitates investigations of systemic insecticides in the laboratory or field (Drinkwater, 1994; Cahill et al., 1996b; Williams et al., 1998). Systemic application can also lead to non-uniform accumulation of insecticide resulting from variable translocation due to
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environmental factors and plant characteristics (Nauen and Elbert, 1994; Palumbo et al., 1996). However, even when uniform coverage is achieved, the properties inherent to the insecticide being studied can alter the behavior of the test arthropods and thereby influence their actual exposure to the toxin (Haynes, 1988; Dombrowski et al., 1994; Longley and Jepson, 1996). Non-uniform coverage of pesticides, when present, makes it extremely difficult to quantify the actual exposure to the test organisms, regardless of the application method. This does not lessen the value of pesticide bioassays using these application methods, because relatively quick and straightforward methods are needed for assessing pesticide effects on beneficial insects. When leaf dip assays are used, effects of uneven coverage can be minimized by use of bioassay chambers with a high proportion of treated area. However, future studies that correlate beneficial insect (or pest) mortality with measurements of toxin on/in the leaf tissue would provide an important step toward quantification of actual exposure viz. insect response. The bioassay method described herein does have limitations. A major constraint is that pesticide application to leaf tissue complicates accurate assessments of exposure to test subjects, especially when leaf dips or systemic applications are used. Also, foliar bioassays are usually not suitable for studies with technical grade insecticides because of volatility and/or phytotoxicity of most organic solvents with which technical products are diluted. For studies using technical insecticides, bioassay methods using glass substrates are advantageous (Plapp, 1971; Cahill and Hackett, 1992). Moreover, foliar bioassays also require space and resources for propagation of high quality plant tissue, and thereby intrinsically require more preparation and setup time than other bioassay methods. In the present study, a low percentage (≤5%) of wasps was captured in the feeding tubes during the course of the exposure period. We do not consider this to be a significant liability to this technique because when working with minute parasitoids it is nearly impossible to completely eliminate entrapment of all wasps in the food source. As discussed earlier, provision of adequate ventilation is a function of the vapor pressure of the insecticides to be studied and the existing air circulation system of the environmental chamber to be used. When necessary, ventilation of the bioassay chambers can be increased by the addition of windows to the chambers or by supplemental fans or pumps (Morse et al., 1986; Hassan and Abdelgader, 2001). However, our results suggest that this effort will not be required in many situations (e.g., as the time interval between application and exposure increases, when tests are conducted below field rates, or when insecticides with low vapor pressures are studied). The limitations of this bioassay procedure not withstanding, it appears to be a valuable tool for investigations of pesticide toxicity to minute Hymenop-
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tera, and may also be adaptable for studies with predators. With necessary modifications, this adaptable bioassay method might have utility in the sequential testing scheme used by the IOBC Working Group on Pesticides and Beneficial Organisms (Sterk et al., 1999). Acknowledgements We thank A.C. Cohen, G. McCain, D.A. Nordlund, and B. Woods (USDAARS BCMRRU, Starkville, MS) for providing A. iole. Technical assistance was provided by M. Sewell and M. Gray. Thanks are extended to C.G. Jackson, G.L. Snodgrass, and anonymous reviewers for their valuable comments on the manuscript. This article reports the results of research only. Mention of a proprietary product does not constitute an endorsement or a recommendation by the USDA for its use. References Bellows, J.T.S. and J.G. Morse, 1993. Toxicity of insecticides used in citrus to Aphytis melinus Debach (Hymenoptera: Aphelinidae) and Rhizobius lophanthae (Blaisd.) (Coleoptera: Coccinellidae). Can. Entomol. 125: 987–994. Bull, D.L. and R.J. Coleman, 1985. Effects of pesticides on Trichogramma spp. Southwest. Entomol. Suppl. 8: 156–168. Cahill, M.R. and B. Hackett, 1992. Insecticidal activity and expression of pyrethroid resistance in adult Bemisia tabaci using a glass vial bioassay. Brighton Crop Protection Conference. pp. 251–256. Cahill, M., I. Denholm, G. Ross, K. Gorman and D. Johnston, 1996a. Relationship between bioassay data and the simulated field performance of insecticides against susceptible and resistant adult Bemisia tabaci (Homoptera: Aleyrodidae). Bull. Entomol. Res. 86: 109– 116. Cahill, M., K. Gorman, S. Day, I. Denholm, A. Elbert and R. Nauen, 1996b. Baseline determination and detection of resistance to imidacloprid in Bemisia tabaci (Homoptera: Aleyrodidae). Bull. Entomol. Res. 86: 343–349. Candolfi, M.P., S. Blümel, R. Forster, F. M. Bakker, C. Grimm, S.A. Hassan, U. Heimbach, M.A. Mead-Briggs, B. Reber, R. Schmuck and G. Vogt (eds), 2000. Guidelines to evaluate side-effects of plant protection products to non-target arthropods. IOBC/WPRS, Gent. Carrillo, J.R., C.G. Jackson, T.D. Carrillo and J. Ellington, 1994. Evaluation of pesticide resistance in Anaphes iole collected from five locations in the western United States. Southwest. Entomol. 19: 157–160. Croft, B.A., 1990. Arthropod Biological Control Agents and Pesticides. John Wiley & Sons, New York. Dittrich, V., 1962. A comparative study of toxicological test methods on a population of the two-spotted spider mite (Tetranychus urticae). J. Econ. Entomol. 55: 644–648. Dombrowski, J.A., S.A. Kolmes and T.J. Dennehy, 1994. Prior exposure to carbaryl alters behavior of Tetranychus urticae Koch on acaricide-treated leaf surfaces. J. Chem. Ecol. 20: 81–90.
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