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Abamectin compatibility with the leafminer parasitoid Diglyphus isaea
Biological Control 35 (2005) 172–179 www.elsevier.com/locate/ybcon
Abamectin compatibility with the leafminer parasitoid Diglyphus isaea Roy Kaspi ¤, Michael P. Parrella Department of Entomology, University of California, One Shields Avenue, Davis, CA 95616, USA Received 25 March 2005; accepted 27 July 2005 Available online 12 September 2005
Abstract The most promising non-chemical approach for controlling Liriomyza leafminers in greenhouses is the augmentative/inoculative releases of the parasitoid Diglyphus isaea. However, the insecticide/miticide abamectin is still an important chemical control method against Liriomyza leafminers and the mite complex that attacks numerous greenhouse crops. It is not uncommon for abamectin to be applied where Diglyphus is being used for leafminer control. Consequently, there is a need to develop compatibility data for this material and Diglyphus. The eVect of abamectin on D. isaea (adults and larvae) was investigated using laboratory and greenhouse experiments. Direct application and uptake of abamectin had a signiWcant negative eVect on the survival of D. isaea adult (both sexes). Abamectin residue on chrysanthemum leaves had a signiWcant negative eVect on adult D. isaea female longevity up to 5 days after application. Abamectin was lethal for D. isaea larvae when applied directly to larvae or when contaminated leafminer larvae were consumed by parasitoid larvae. However, the percent emergence of D. isaea was not aVected by abamectin treatments when applied to chrysanthemum plants that contained parasitoid larvae. Moreover, the longevity of these emerged adults was not aVected by abamectin applications. In light of these data, abamectin compatibility with D. isaea for leafminer control in IPM programs for greenhouses is discussed. 2005 Elsevier Inc. All rights reserved. Keywords: Diglyphus isaea; Liriomyza trifolii; Abamectin compatibility; Biological control; IPM; Greenhouse
1. Introduction The leafminer Liriomyza trifolii (Burgess) is a worldwide pest of ornamental and vegetable crops. L. trifolii is a polyphagous leafminer (ca. 400 diVerent host plants; Murphy and LaSalle, 1999; Parrella, 1987) that undergoes larval development in the plant leaf tissue and forms serpentine mines within the leaves (Parrella and Bethke, 1988). Damage is caused mostly by larvae that feed their way inside the plant-host mesophyll, and by the female feeding behavior, (puncturing the leaf with their ovipositor, and feeding from the leaf sap), an action that decreases the plant’s photosynthesis (Parrella et al.,
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1985), provides entry sites for plant pathogens (Broadbent and Matteoni, 1990; Matteoni and Broadbent, 1988), and creates small marks that reduces the aesthetic appearance of leaves. The translaminar pesticide, abamectin, has been a primary means of controlling Liriomyza leafminers. Abamectin is a streptomycete derived agonist of -aminobutyric acid (Burg et al., 1979; Putter et al., 1981), and is very eVective against leafmining Agromyzidae (Cox et al., 1995; Hara, 1986; Leibee, 1988; Parrella et al., 1988; Trumble, 1985; Weintraub, 1999, 2001). In addition to leafminer control, abamectin is used for controlling mites, thrips, aphids, whiteXies, psyllids, diaspid scale insects, and lepidopteran pest species (Lasota and Dybas, 1991). The most promising non-chemical approach for controlling Liriomyza leafminers in greenhouses is the
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augmentative releases of the oligophagous larval ectoparasitoid D. isaea (Walker) (Hymenoptera: Eulophidae) (e.g., Boot et al., 1992; Minkenberg and Van Lenteren, 1987; Ozawa et al., 2001). D. isaea is a facultative gregarious parasitoid and the adult females lay one to Wve eggs adjacent to late leafminer larval stages (Hendrickson, 1975; Minkenberg and Van Lenteren, 1986, 1987). The parasitoid larvae feed on the host for ca. 5 days (at 25 °C, on L. trifolii), then pupate in the mine. The adult parasitoids emerge after about 5.5 days (at 25 °C, on L. trifolii from round holes that they cut in the leaves) and live for about 10 days (Minkenberg, 1989). Integrated control was Wrst deWned as “applied pest control that combines and integrates biological and chemical control. Chemical control is used as necessary and in a manner which is least disruptive to biological control. Integrated control may make use of naturally occurring biological control as well as biological control aVected by manipulated or induced biotic agents” (Stern et al., 1959). This early deWnition still is true today (Kogan, 1998). An ideal pesticide for incorporation into a greenhouse Integrated Pest Management (IPM) program is pest speciWc and not harmful to the pest’s biological agents (e.g., Hoy, 2000; Hoy and Ouyang, 1989), or has a short residual eVect on the pest’s natural enemies. In this case, the residual toxicity period should be taken into consideration prior to releasing biological agents following a pesticide application (Malezieux et al., 1992; Shipp et al., 2000). Because abamectin and the leafminer parasitoid D. isaea are key methods in combating leafminers in greenhouses, and the use of abamectin is also important for the control of other major greenhouse pests such as spider mites, the objective of our study was to evaluate the toxicity of abamectin to D. isaea (larvae and adults). The following research questions were addressed: (1) will abamectin kill adult D. isaea via direct contact? (2) will abamectin kill D. isaea if adults feed on contaminated honey solutions? (3) will abamectin kill adult D. isaea through residual action, and if so, for how long? (4) will abamectin kill D. isaea larvae if they are sprayed while protected in the leaf? (5) will abamectin impact the longevity of adult D. isaea if they are sprayed as larvae in the leaf? (6) will abamectin kill D. isaea larvae if there is direct contact with the larvae? and (7) will abamectinpoisoned leafminer larvae kill D. isaea larvae feeding on them?
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tems,’ Berkel en Rodenrijs, The Netherlands) were used within a few days of receipt (they had been refrigerated at 10 § 1 °C until then), and the pesticide abamectin (‘Avid 0.15 EC’ Syngenta Crop Protection, Greensboro, NC) was used at the commercial rate (0.62 ml/liter) recommended for leafminer control. 2.1. EVect of direct abamectin application on adult Diglyphus Diglyphus males and females were placed individually in small containers (18 § 2 mm high, 22 mm diameter, plastic cylinder netted on the top and bottom). Using an air-brush sprayer (3000S Airbrush Kit, Aztec, Rockford, IL, USA) the containers were sprayed to drip with 0.5 ml water (nfemale D 56, nmale D 36), abamectin (nfemale D 60, nmale D 36), or were not sprayed [untreated control (UTC); nfemale D 33, nmale D 28]. After application, the cages were placed randomly in a growth chamber (14:10 L:D, 70 § 20% r.h. and at 25 § 1 °C), where the parasitoids had ad libitum access to diluted honey (1:1 honey/ water). Diglyphus mortality was recorded daily for 50 days. To evaluate the eVects of abamectin on Diglyphus longevity, Kaplan–Meier survival analyses were used (both Log-rank (weights later events) and Wilcoxon (weights early events) tests; SAS Institute, 2005). Planned multiple comparisons were performed using Kaplan–Meier survival analyses with an adjustment of the Bonferroni procedure (Holm, 1979). Adjusted (⬘) was noted in the results only when it diVered from D 0.05. 2.2. EVect of abamectin intake on adult Diglyphus Male and female Diglyphus were starved for 24 h and then placed individually in Petri dishes (55 mm diameter) that contained a drop of honey and water (1:1) (nfemale D 19, nmale D 18) or honey and abamectin solution (1:1) (nfemale D 25, nmale D 22). The feeding time for each parasitoid was recorded. After feeding ceased, each parasitoid was placed individually into small containers that were positioned randomly in a growth chamber (conditions described earlier), where they had ad libitum access to diluted honey. Diglyphus mortality was recorded daily for 25 days. To investigate the eVects of abamectin and feeding duration on Diglyphus longevity, proportional hazard survival analyses were performed and signiWcance was tested using likelihood ratio tests. Non-signiWcant (P > 0.05) interactions were discarded in Wnal models (SAS Institute, 2005).
2. Materials and methods Experiments were carried out in a laboratory and greenhouse located on the campus of the University of California, Davis. In all experiments, commercial D. isaea parasitoids (‘Miglyphus’, ‘Koppert-Biological Sys-
2.3. EVect of abamectin leaf residue on adult female Diglyphus Chrysanthemum plants (var. ‘Miramar’) were sprayed to drip with water (n D 3 per experiment) or abamectin
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(n D 3 per experiment) using a hand sprayer. One, 5, 7, and 10 days later, small netted containers (20 § 0.5 mm high, 11 mm diameter, plastic cylinder netted on top) were fastened to individual leaves with clips (MelamedMadjar et al., 1984). One Diglyphus female was placed in each container. Water was smeared on the container nets on a daily basis. The plants were placed in a random order inside a large cage (47 £ 47 £ 43 cm, with a glass on top) in the laboratory (16:8 L:D, 53 § 25% r.h. and at 25.5 § 5 °C). Diglyphus mortality was recorded daily for the next 15 days. To evaluate the eVects of abamectin on Diglyphus longevity, Kaplan–Meier survival analyses were used (both Log-rank and Wilcoxon tests; SAS Institute, 2005). 2.4. EVect of abamectin application on Diglyphus larvae in the leaves The experiment was conducted in October–November 2003 (i.e., ‘Block I’; nabamectin D 19; nwater D 18) and April–May 2004 (i.e., ‘Block II’; nabamectin D 25; nwater D 25), in a greenhouse located on the University of California, Davis campus. Three female and two male leafminers (1- to 5-day old) were placed in a ‘screened cage’ (25 § 0.5 cm high £ 15.2 cm diameter), with a chrysanthemum plant that was pinched after 4–5 weeks of growth. The caged pots were placed randomly on greenhouse “ebb & Xood” tables (Agro Dynamics, East Brunswick, NJ; Block I: 23.5 § 6.5 °C, 59 § 25% r.h., 14:10 L:D, Block II: 22 § 11 °C, 59 § 40% r.h., 14:10 L:D). Three days later, the Xies were removed from all of the cages. When second and third instar larvae were detected (5–6 days later), two females and one male Diglyphus were released into all cages, and diluted honey was streaked on the top of the cages. Three days later, the parasitoids were removed from all cages. After three more days (when Diglyphus larvae were present) the plants were sprayed to drip with water or abamectin using a backpack sprayer (RL FLO-MASTER, model: 2603HD, Lowell, MI). A week later the plants were cut at the soil line and individually placed inside 940 ml glass containers (netted on top) located in the laboratory (23 § 2 °C, 40 § 15% r.h., 16:8 L:D). Diglyphus adults were allowed to emerge freely, and were counted after all died. To compare the eVect of abamectin application on Diglyphus larvae in the leaves, data were square root transformed and subjected to a two-way analysis of variance (ANOVA), using block and treatment as model factors. Data are reported as means § SE. 2.5. EVect of abamectin applied to Diglyphus larvae in leaves on adult longevity In the laboratory, six chrysanthemum plants were placed together inside a large cage (47 £ 47 £ 43 cm) with a glass top. Approximately 500 leafminers were released
into the cage for 24 h, and then removed. Six days later when second and third instar larvae were detected, approximately 250 Diglyphus adults (ca. 1:1 sex ratio) were released into the cage and removed after 24 h. Three days after the plants were exposed to the Diglyphus adults, the plants were sprayed to drip with water (three plants) or abamectin (three plants). Four days later, Diglyphus pupae were removed from their mines (using an insect pin #2 and a Wne brush #00) and placed individually in Petri dishes (60 £ 15 mm), Wlled with 0.5 mm layers of Agar (1%) that was covered with Wlter paper (55 mm; Whatman International, Maidstone, England). The Petri dishes (nabamectin D 38; nwater D 27) were placed randomly inside a growth chamber (25 § 1 °C, 60 § 20% r.h., 14:10 L:D). The Petri dishes were inspected daily for 1 week, and the adult emergence percentage and longevity were recorded, and evaluated using Fisher’s exact test (Zar, 1999) and Kaplan–Meier survival analyses (SAS Institute, 2005). 2.6. EVect of direct abamectin contact with Diglyphus larvae Forty-seven chrysanthemum leaves infested by Diglyphus larvae (second and third instars) were removed from their plants. While viewing under a stereoscopic microscope, the mines were opened and Diglyphus larvae were touched with abamectin (n D 19), water (n D 18), or were left untouched (‘UTC’, n D 10) using a Wne brush (#00). The mines were then reclosed, and the leaves were placed in Petri dishes (with agar as a moisture source covered with Wlter paper, as described in Section 2.5). The Petri dishes were placed in a growth chamber (as described in Section 2.5) and the number of emerged adults was recorded. To compare the proportion of adult emergence, Pearson 2 test was used (Zar, 1999). Pairwise comparisons were performed using multiple Fisher’s exact tests with an adjustment of the classical Bonferroni procedure (Holm, 1979). Adjusted (⬘) was noted in the results only when it diVered from D 0.05. 2.7. EVect of abamectin-poisoned leafminer larvae on Diglyphus larvae Leafminer and Diglyphus larvae were obtained as described in Section 2.5. Three chrysanthemum plants were sprayed to drip with water or abamectin. Twentyfour hours later, leaves that contained actively feeding leafminer larvae from both treated and untreated plants were put in a freezer (¡16 § 4 °C) for 15 min. Forty-one chrysanthemum leaves infested with Diglyphus larvae (second and third instars) were cut from their plants. While viewing under a stereoscopic microscope, the mines were opened and the leafminer larvae were replaced with the frozen larvae that consumed abamectin (n D 22) or water (n D 19) treated leaves. Then the
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P < 0.0001; Wilcoxon test: 2 D 63.000, df D 1, P < 0.0001; ⬘ D 0.0166). In addition, males that were treated with water suVered a higher mortality rate than that of the untreated males (Log-rank test: 2 D 9.975, df D 1, P D 0.0016; Wilcoxon test: 2 D 14.8399, df D 1, P < 0.0001; ⬘ D 0.0166).
mines were closed back, and the leaves were placed in Petri dishes (described earlier). The Petri dishes were placed in a growth chamber, and the number of adult Diglyphus emerging was recorded. To compare the proportion of adult emergence, Fisher’s exact test was used (Zar, 1999).
3.2. EVect of abamectin intake on adult Diglyphus 3. Results Feeding duration did not aVect Diglyphus survival (female: 2 < 0.001, df D 1, P D 0.9889; male: 2 D 0.013, df D 1, P D 0.9108). However, abamectin intake signiWcantly aVected Diglyphus survival (females: 2 D 32.482, df D 1, P < 0.0001; males: 2 D 29.243, df D 1, P < 0.0001). Abamectin intake (both by females and males) severely increased mortality rate (Fig. 2). No interaction was detected between feeding treatments (abamectin-treated honey or honey only) and feeding durations (female: 2 < 0.001, df D 1, P D 1.00; male: 2 D 0.759, df D 1, P D 0.3835).
3.1. EVect of direct abamectin application on adult Diglyphus We found that abamectin treatment had a signiWcant negative eVect on female Diglyphus survival (Log-rank test: 2 D 148.000, df D 2, P < 0.0001; Wilcoxon test: 2 D 148.008, df D 2, P < 0.0001; Fig. 1A). Females that were treated with direct abamectin application, died within 24 h of treatment. This mortality rate was signiWcantly higher than that of the water treated females (Log-rank test: 2 D 115.000, df D 1, P < 0.0001; Wilcoxon test: 2 D 115.000, df D 1, P < 0.0001; ⬘ D 0.0166) or the untreated females (Log-rank test: 2 D 92.000, df D 1, P < 0.0001; Wilcoxon test: 2 D 92.000, df D 1, P < 0.0001; ⬘ D 0.0166). In contrast, there was no diVerence between water treated and untreated female survival (Log-rank test: 2 D 0.001, df D 1, P D 0.9743; Wilcoxon test: 2 D 0.029, df D 1, P D 0.8644; ⬘ D 0.0166). Similar to female survival, the abamectin treatment had a signiWcant negative eVect on male Diglyphus survival (Logrank test: 2 D 94.964, df D 2, P < 0.0001; Wilcoxon test: 2 D 93.731, df D 2, P < 0.0001; Fig. 1B). Males that were treated directly with abamectin died within 24 h of treatment. This mortality rate was signiWcantly higher than that of the water treated males (Log-rank test: 2 D 60.077, df D 1, P < 0.0001; Wilcoxon test: 2 D 60.077, df D 1, P < 0.0001; ⬘ D 0.0166) or the untreated males (Log-rank test: 2 D 63.000, df D 1, A
3.3. EVect of abamectin leaf residue on adult female Diglyphus When female Diglyphus were conWned within clipcages to chrysanthemum leaves that were treated with abamectin or water 24 h after application, abamectin had a signiWcant negative eVect on D. isaea adult female longevity (Log-rank test: 2 D 7.333, df D 1, P D 0.0068; Wilcoxon test: 2 D 12.971, df D 1, P D 0.0003; nabamectin D 22, nwater D 26; Fig. 3A). Although this was reduced Wve days after application, abamectin leaf residue still had a signiWcant eVect on female longevity (Log-rank test: 2 D 3.622, df D 1, P D 0.0570; Wilcoxon test: 2 D 3.569, df D 1, P D 0.0588; nabamectin D 24, nwater D 23; Fig. 3B). Longevity of females that were conWned to leaves seven (Log-rank test: 2 D 0.057, df D 1, P D 0.8117; Wilcoxon test: 2 D 0.007, df D 1, P D 0.9326;
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Fig. 1. EVect of direct abamectin spray on adult D. isaea; lines show Kaplan–Meier estimates of the probability of (A) female, and (B) male survival. DiVerent letters indicate a signiWcant diVerence (P < 0.0166). ‘UTC’ D untreated control.
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Fig. 2. EVect of abamectin intake on adult D. isaea. Lines show Kaplan–Meier estimates of the probability of (A) female, and (B) male survival. DiVerent letters indicate a signiWcant diVerence (P < 0.05).
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Fig. 3. EVect of leaf residue on adult female D. isaea longevity. (A) Residual toxicity eVect one day after application. (B) Residual toxicity eVect Wve days after application. (C) Residual toxicity eVect seven days after application. (D) Residual toxicity eVect 10 days after application.
nabamectin D 26, nwater D 29; Fig. 3C) and 10 days after application was not aVected by abamectin treatments (Log-rank test: 2 D 2.067, df D 1, P D 0.1505; Wilcoxon test: 2 D 0.276, df D 1, P D 0.5992; nabamectin D 25, nwater D 22; Fig. 3D).
3.4. EVect of abamectin application on Diglyphus larvae in the leaves Although Diglyphus emergence varied between blocks (F D 25.977, df D 1, P < 0.0001) under greenhouse condi-
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Fig. 4. EVect of abamectin application on emergence of D. isaea adults from chrysanthemum plants in the greenhouse. Horizontal lines denote the mean (§SE) and diamonds 95% conWdence intervals.
tions, abamectin application did not signiWcantly aVect the numbers of emerged adult Diglyphus (F D 0.496, df D 1, P D 0.4834; Fig. 4). No interaction was detected between blocks and treatments (F D 2.052, df D 1, P D 0.1558). 3.5. EVect of abamectin applied to Diglyphus larvae in leaves on adult longevity When applied to leaves containing Diglyphus larvae, abamectin did not have a signiWcant eVect on the percentage of Diglyphus emerging (abamectin: 84.21% emerged; water: 92.59% emerged; P D 0.2692) or on the longevity of emerged Diglyphus adults (Log-rank test: 2 D 0.007, df D 1, P D 0.9309; Wilcoxon test: 2 D 0.1367, df D 1, P D 0.7116; Fig. 5). 3.6. EVect of direct abamectin contact with Diglyphus larvae We found that direct larval contact with abamectin had a signiWcant negative eVect on Diglyphus adult emergence (2 D 20.579, df D 2, P < 0.0001); 100% of abamectin treated Diglyphus did not survive (Fig. 6). Abamectin application signiWcantly aVected Diglyphus survival compared to the water treatment (P D 0.0004, ⬘ D 0.0166) and with the UTC treatment (P < 0.0001, ⬘ D 0.0166). No diVerence was found between the water and UTC treatments (P D 0.1237, ⬘ D 0.0166). 3.7. EVect of abamectin-poisoned leafminer larvae on Diglyphus larvae When untreated Diglyphus host larvae were replaced with larvae that were taken from plants treated with
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Fig. 6. EVect of direct abamectin contact with D. isaea larvae on the percentage of adult emergence. ‘UTC’ D untreated control. DiVerent letters indicate a signiWcant diVerence (P < 0.0166).
water or abamectin, abamectin-poisoned leafminer larvae had a signiWcant negative eVect on Diglyphus larvae survival (P < 0.0001); 91% of those larvae did not survive (Fig. 7).
4. Discussion Previous studies on the eVect of abamectin on Diglyphus spp. are confusing and conXicting. Abamectin was reported to have a “minimal eVect” on Diglyphus intermedius (Girault) and Diglyphus begini (Ashmead) in celery Welds (Trumble, 1985). Similarly, Hara (1986) found no abamectin adverse eVect on immature D. intermedius stages in cut chrysanthemum crops. In this study Hara (1986) found that D. isaea accounted for less than one percent of the total recovered parasitoid species, but no data were presented on D. isaea recov-
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Percentage of adult emergence
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Fig. 7. EVect of abamectin-poisoned leafminer larvae on D. isaea emergence percentage. DiVerent letters indicate a signiWcant diVerence (P < 0.05).
ery in the control vs. treated plots. In a celery Weld study, Weintraub (1999) showed that abamectin (plus 1% UltraWne oil) applications did not aVect the D. isaea adult population. In contrast, Schuster (1994) used laboratory tomato leaf-dip experiments and found that abamectin causes 73% D. intermedius larval mortality, and 38% D. intermedius pupa mortality. In a potato Weld study, Weintraub (2001) showed a reduction in the number of D. isaea immature stages only 14 days after a sole abamectin application. This D. isaea immature population started to recover 28 days from application and by the end of the study equaled the D. isaea immature population in the control plots. Prijono et al. (2004) and Schuster (1994) showed (in laboratory studies) that abamectin residue causes some mortality of D. isaea adults when they were exposed immediately after application. Our Wndings show that abamectin is lethal to the adult D. isaea (both sexes) when direct contact is made, meaning that an abamectin application in a greenhouse would eliminate most if not all D. isaea adults present. Moreover, adults that are released or emerge a few days later, and feed on food contaminated with abamectin (such as contaminated water or honeydew) would also be negatively aVected. Furthermore, parasitoid longevity would be negatively aVected if they contact recently (1–5 days) sprayed leaves. These results support previous studies that showed negative aVects of abamectin residue on Diglyhus (Schuster, 1994; Prijono et al., 2004). However, our results suggest that the abamectin residues were depleted 7 days after application under laboratory conditions and therefore a period of one week is suggested as the duration of time before releases of D. isaea into abamectin treated greenhouses can be made. Under Weld conditions, abamectin leaf residues can rapidly degrade (half-life < one day) probably caused by photodegradation (Bull et al., 1984). Greenhouses vastly vary in the extent to which
they block ultraviolet light. Therefore, abamectin residue depletion period may be shorter under some greenhouse conditions. These Wndings are similar to those of another leafminer parasitoid—Dacnusa sibirica (Shipp et al., 2000), where under cucumber greenhouse conditions adult mortality was less than 10% when they were exposed to abamectin treated leaves 6 days after application. Our results also show that abamectin is very toxic to D. isaea larvae when direct contact is made or when they feed on abamectin-poisoned leafminer larvae. Both of these experiments were done in highly manipulated laboratory experiments. These types of experiments were necessary to better understand the mechanism by which Diglyphus avoids abamectin poisoning. While they are inside the mine D. isaea larvae are protected and are not aVected by abamectin application (both under greenhouse and laboratory conditions). Abamectin probably does not penetrate the dead plant tissue and there is air surrounding both the parasitoid and its leafminer host. In addition, the hosts of immature D. isaea (i.e., leafminer larvae) are paralyzed or dead at this point, so they do not consume abamectin. Therefore, physical aspects of the mines and the fact that Diglyphus paralyzes its host provides an ecological refuge for D. isaea larvae from abamactin applications. Moreover, about 6–8 days after application (at 25 °C), adults should emerge into a relatively safe environment in terms of abamectin leaf residue. Abamectin application did not have a signiWcant eVect on D. isaea longevity after emerging from treated chrysanthemum plants. However, abamectin may cause some sub-lethal impacts on D. isaea (other than longevity) such as abnormal behavior, as well as decreased fecundity, health, and Wtness. These possible sub-lethal impacts of abamectin on D. isaea were not evaluated in this study. Despite being an excellent leafminer insecticide, our results suggest that abamectin is compatible with the parasitoid D. isaea for leafminer control in greenhouses provided that the duration of its residual toxicity is taken into account.
Acknowledgments We thank Thomas Costamagna, Bob Starnes, and Trinity Strehl for their invaluable advice and assistance to our study. We acknowledge the generous donation of Chrysanthemum cuttings by ‘Yoder Brothers,’ yellow sticky cards by ‘Seabright Laboratories,’ and Diglyphus by ‘Plant Sciences, Watsonville, CA, and ‘Koppert-Biological Systems.’ This research was supported by the National Research Initiative (NRI; USDA) Competitive Grants program, Proposal No. 2002-01971; and the American Floral Endowment.
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Abamectin compatibility with the leafminer parasitoid Diglyphus isaea Roy Kaspi ¤, Michael P. Parrella Department of Entomology, University of California, One Shields Avenue, Davis, CA 95616, USA Received 25 March 2005; accepted 27 July 2005 Available online 12 September 2005
Abstract The most promising non-chemical approach for controlling Liriomyza leafminers in greenhouses is the augmentative/inoculative releases of the parasitoid Diglyphus isaea. However, the insecticide/miticide abamectin is still an important chemical control method against Liriomyza leafminers and the mite complex that attacks numerous greenhouse crops. It is not uncommon for abamectin to be applied where Diglyphus is being used for leafminer control. Consequently, there is a need to develop compatibility data for this material and Diglyphus. The eVect of abamectin on D. isaea (adults and larvae) was investigated using laboratory and greenhouse experiments. Direct application and uptake of abamectin had a signiWcant negative eVect on the survival of D. isaea adult (both sexes). Abamectin residue on chrysanthemum leaves had a signiWcant negative eVect on adult D. isaea female longevity up to 5 days after application. Abamectin was lethal for D. isaea larvae when applied directly to larvae or when contaminated leafminer larvae were consumed by parasitoid larvae. However, the percent emergence of D. isaea was not aVected by abamectin treatments when applied to chrysanthemum plants that contained parasitoid larvae. Moreover, the longevity of these emerged adults was not aVected by abamectin applications. In light of these data, abamectin compatibility with D. isaea for leafminer control in IPM programs for greenhouses is discussed. 2005 Elsevier Inc. All rights reserved. Keywords: Diglyphus isaea; Liriomyza trifolii; Abamectin compatibility; Biological control; IPM; Greenhouse
1. Introduction The leafminer Liriomyza trifolii (Burgess) is a worldwide pest of ornamental and vegetable crops. L. trifolii is a polyphagous leafminer (ca. 400 diVerent host plants; Murphy and LaSalle, 1999; Parrella, 1987) that undergoes larval development in the plant leaf tissue and forms serpentine mines within the leaves (Parrella and Bethke, 1988). Damage is caused mostly by larvae that feed their way inside the plant-host mesophyll, and by the female feeding behavior, (puncturing the leaf with their ovipositor, and feeding from the leaf sap), an action that decreases the plant’s photosynthesis (Parrella et al.,
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1985), provides entry sites for plant pathogens (Broadbent and Matteoni, 1990; Matteoni and Broadbent, 1988), and creates small marks that reduces the aesthetic appearance of leaves. The translaminar pesticide, abamectin, has been a primary means of controlling Liriomyza leafminers. Abamectin is a streptomycete derived agonist of -aminobutyric acid (Burg et al., 1979; Putter et al., 1981), and is very eVective against leafmining Agromyzidae (Cox et al., 1995; Hara, 1986; Leibee, 1988; Parrella et al., 1988; Trumble, 1985; Weintraub, 1999, 2001). In addition to leafminer control, abamectin is used for controlling mites, thrips, aphids, whiteXies, psyllids, diaspid scale insects, and lepidopteran pest species (Lasota and Dybas, 1991). The most promising non-chemical approach for controlling Liriomyza leafminers in greenhouses is the
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augmentative releases of the oligophagous larval ectoparasitoid D. isaea (Walker) (Hymenoptera: Eulophidae) (e.g., Boot et al., 1992; Minkenberg and Van Lenteren, 1987; Ozawa et al., 2001). D. isaea is a facultative gregarious parasitoid and the adult females lay one to Wve eggs adjacent to late leafminer larval stages (Hendrickson, 1975; Minkenberg and Van Lenteren, 1986, 1987). The parasitoid larvae feed on the host for ca. 5 days (at 25 °C, on L. trifolii), then pupate in the mine. The adult parasitoids emerge after about 5.5 days (at 25 °C, on L. trifolii from round holes that they cut in the leaves) and live for about 10 days (Minkenberg, 1989). Integrated control was Wrst deWned as “applied pest control that combines and integrates biological and chemical control. Chemical control is used as necessary and in a manner which is least disruptive to biological control. Integrated control may make use of naturally occurring biological control as well as biological control aVected by manipulated or induced biotic agents” (Stern et al., 1959). This early deWnition still is true today (Kogan, 1998). An ideal pesticide for incorporation into a greenhouse Integrated Pest Management (IPM) program is pest speciWc and not harmful to the pest’s biological agents (e.g., Hoy, 2000; Hoy and Ouyang, 1989), or has a short residual eVect on the pest’s natural enemies. In this case, the residual toxicity period should be taken into consideration prior to releasing biological agents following a pesticide application (Malezieux et al., 1992; Shipp et al., 2000). Because abamectin and the leafminer parasitoid D. isaea are key methods in combating leafminers in greenhouses, and the use of abamectin is also important for the control of other major greenhouse pests such as spider mites, the objective of our study was to evaluate the toxicity of abamectin to D. isaea (larvae and adults). The following research questions were addressed: (1) will abamectin kill adult D. isaea via direct contact? (2) will abamectin kill D. isaea if adults feed on contaminated honey solutions? (3) will abamectin kill adult D. isaea through residual action, and if so, for how long? (4) will abamectin kill D. isaea larvae if they are sprayed while protected in the leaf? (5) will abamectin impact the longevity of adult D. isaea if they are sprayed as larvae in the leaf? (6) will abamectin kill D. isaea larvae if there is direct contact with the larvae? and (7) will abamectinpoisoned leafminer larvae kill D. isaea larvae feeding on them?
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tems,’ Berkel en Rodenrijs, The Netherlands) were used within a few days of receipt (they had been refrigerated at 10 § 1 °C until then), and the pesticide abamectin (‘Avid 0.15 EC’ Syngenta Crop Protection, Greensboro, NC) was used at the commercial rate (0.62 ml/liter) recommended for leafminer control. 2.1. EVect of direct abamectin application on adult Diglyphus Diglyphus males and females were placed individually in small containers (18 § 2 mm high, 22 mm diameter, plastic cylinder netted on the top and bottom). Using an air-brush sprayer (3000S Airbrush Kit, Aztec, Rockford, IL, USA) the containers were sprayed to drip with 0.5 ml water (nfemale D 56, nmale D 36), abamectin (nfemale D 60, nmale D 36), or were not sprayed [untreated control (UTC); nfemale D 33, nmale D 28]. After application, the cages were placed randomly in a growth chamber (14:10 L:D, 70 § 20% r.h. and at 25 § 1 °C), where the parasitoids had ad libitum access to diluted honey (1:1 honey/ water). Diglyphus mortality was recorded daily for 50 days. To evaluate the eVects of abamectin on Diglyphus longevity, Kaplan–Meier survival analyses were used (both Log-rank (weights later events) and Wilcoxon (weights early events) tests; SAS Institute, 2005). Planned multiple comparisons were performed using Kaplan–Meier survival analyses with an adjustment of the Bonferroni procedure (Holm, 1979). Adjusted (⬘) was noted in the results only when it diVered from D 0.05. 2.2. EVect of abamectin intake on adult Diglyphus Male and female Diglyphus were starved for 24 h and then placed individually in Petri dishes (55 mm diameter) that contained a drop of honey and water (1:1) (nfemale D 19, nmale D 18) or honey and abamectin solution (1:1) (nfemale D 25, nmale D 22). The feeding time for each parasitoid was recorded. After feeding ceased, each parasitoid was placed individually into small containers that were positioned randomly in a growth chamber (conditions described earlier), where they had ad libitum access to diluted honey. Diglyphus mortality was recorded daily for 25 days. To investigate the eVects of abamectin and feeding duration on Diglyphus longevity, proportional hazard survival analyses were performed and signiWcance was tested using likelihood ratio tests. Non-signiWcant (P > 0.05) interactions were discarded in Wnal models (SAS Institute, 2005).
2. Materials and methods Experiments were carried out in a laboratory and greenhouse located on the campus of the University of California, Davis. In all experiments, commercial D. isaea parasitoids (‘Miglyphus’, ‘Koppert-Biological Sys-
2.3. EVect of abamectin leaf residue on adult female Diglyphus Chrysanthemum plants (var. ‘Miramar’) were sprayed to drip with water (n D 3 per experiment) or abamectin
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(n D 3 per experiment) using a hand sprayer. One, 5, 7, and 10 days later, small netted containers (20 § 0.5 mm high, 11 mm diameter, plastic cylinder netted on top) were fastened to individual leaves with clips (MelamedMadjar et al., 1984). One Diglyphus female was placed in each container. Water was smeared on the container nets on a daily basis. The plants were placed in a random order inside a large cage (47 £ 47 £ 43 cm, with a glass on top) in the laboratory (16:8 L:D, 53 § 25% r.h. and at 25.5 § 5 °C). Diglyphus mortality was recorded daily for the next 15 days. To evaluate the eVects of abamectin on Diglyphus longevity, Kaplan–Meier survival analyses were used (both Log-rank and Wilcoxon tests; SAS Institute, 2005). 2.4. EVect of abamectin application on Diglyphus larvae in the leaves The experiment was conducted in October–November 2003 (i.e., ‘Block I’; nabamectin D 19; nwater D 18) and April–May 2004 (i.e., ‘Block II’; nabamectin D 25; nwater D 25), in a greenhouse located on the University of California, Davis campus. Three female and two male leafminers (1- to 5-day old) were placed in a ‘screened cage’ (25 § 0.5 cm high £ 15.2 cm diameter), with a chrysanthemum plant that was pinched after 4–5 weeks of growth. The caged pots were placed randomly on greenhouse “ebb & Xood” tables (Agro Dynamics, East Brunswick, NJ; Block I: 23.5 § 6.5 °C, 59 § 25% r.h., 14:10 L:D, Block II: 22 § 11 °C, 59 § 40% r.h., 14:10 L:D). Three days later, the Xies were removed from all of the cages. When second and third instar larvae were detected (5–6 days later), two females and one male Diglyphus were released into all cages, and diluted honey was streaked on the top of the cages. Three days later, the parasitoids were removed from all cages. After three more days (when Diglyphus larvae were present) the plants were sprayed to drip with water or abamectin using a backpack sprayer (RL FLO-MASTER, model: 2603HD, Lowell, MI). A week later the plants were cut at the soil line and individually placed inside 940 ml glass containers (netted on top) located in the laboratory (23 § 2 °C, 40 § 15% r.h., 16:8 L:D). Diglyphus adults were allowed to emerge freely, and were counted after all died. To compare the eVect of abamectin application on Diglyphus larvae in the leaves, data were square root transformed and subjected to a two-way analysis of variance (ANOVA), using block and treatment as model factors. Data are reported as means § SE. 2.5. EVect of abamectin applied to Diglyphus larvae in leaves on adult longevity In the laboratory, six chrysanthemum plants were placed together inside a large cage (47 £ 47 £ 43 cm) with a glass top. Approximately 500 leafminers were released
into the cage for 24 h, and then removed. Six days later when second and third instar larvae were detected, approximately 250 Diglyphus adults (ca. 1:1 sex ratio) were released into the cage and removed after 24 h. Three days after the plants were exposed to the Diglyphus adults, the plants were sprayed to drip with water (three plants) or abamectin (three plants). Four days later, Diglyphus pupae were removed from their mines (using an insect pin #2 and a Wne brush #00) and placed individually in Petri dishes (60 £ 15 mm), Wlled with 0.5 mm layers of Agar (1%) that was covered with Wlter paper (55 mm; Whatman International, Maidstone, England). The Petri dishes (nabamectin D 38; nwater D 27) were placed randomly inside a growth chamber (25 § 1 °C, 60 § 20% r.h., 14:10 L:D). The Petri dishes were inspected daily for 1 week, and the adult emergence percentage and longevity were recorded, and evaluated using Fisher’s exact test (Zar, 1999) and Kaplan–Meier survival analyses (SAS Institute, 2005). 2.6. EVect of direct abamectin contact with Diglyphus larvae Forty-seven chrysanthemum leaves infested by Diglyphus larvae (second and third instars) were removed from their plants. While viewing under a stereoscopic microscope, the mines were opened and Diglyphus larvae were touched with abamectin (n D 19), water (n D 18), or were left untouched (‘UTC’, n D 10) using a Wne brush (#00). The mines were then reclosed, and the leaves were placed in Petri dishes (with agar as a moisture source covered with Wlter paper, as described in Section 2.5). The Petri dishes were placed in a growth chamber (as described in Section 2.5) and the number of emerged adults was recorded. To compare the proportion of adult emergence, Pearson 2 test was used (Zar, 1999). Pairwise comparisons were performed using multiple Fisher’s exact tests with an adjustment of the classical Bonferroni procedure (Holm, 1979). Adjusted (⬘) was noted in the results only when it diVered from D 0.05. 2.7. EVect of abamectin-poisoned leafminer larvae on Diglyphus larvae Leafminer and Diglyphus larvae were obtained as described in Section 2.5. Three chrysanthemum plants were sprayed to drip with water or abamectin. Twentyfour hours later, leaves that contained actively feeding leafminer larvae from both treated and untreated plants were put in a freezer (¡16 § 4 °C) for 15 min. Forty-one chrysanthemum leaves infested with Diglyphus larvae (second and third instars) were cut from their plants. While viewing under a stereoscopic microscope, the mines were opened and the leafminer larvae were replaced with the frozen larvae that consumed abamectin (n D 22) or water (n D 19) treated leaves. Then the
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P < 0.0001; Wilcoxon test: 2 D 63.000, df D 1, P < 0.0001; ⬘ D 0.0166). In addition, males that were treated with water suVered a higher mortality rate than that of the untreated males (Log-rank test: 2 D 9.975, df D 1, P D 0.0016; Wilcoxon test: 2 D 14.8399, df D 1, P < 0.0001; ⬘ D 0.0166).
mines were closed back, and the leaves were placed in Petri dishes (described earlier). The Petri dishes were placed in a growth chamber, and the number of adult Diglyphus emerging was recorded. To compare the proportion of adult emergence, Fisher’s exact test was used (Zar, 1999).
3.2. EVect of abamectin intake on adult Diglyphus 3. Results Feeding duration did not aVect Diglyphus survival (female: 2 < 0.001, df D 1, P D 0.9889; male: 2 D 0.013, df D 1, P D 0.9108). However, abamectin intake signiWcantly aVected Diglyphus survival (females: 2 D 32.482, df D 1, P < 0.0001; males: 2 D 29.243, df D 1, P < 0.0001). Abamectin intake (both by females and males) severely increased mortality rate (Fig. 2). No interaction was detected between feeding treatments (abamectin-treated honey or honey only) and feeding durations (female: 2 < 0.001, df D 1, P D 1.00; male: 2 D 0.759, df D 1, P D 0.3835).
3.1. EVect of direct abamectin application on adult Diglyphus We found that abamectin treatment had a signiWcant negative eVect on female Diglyphus survival (Log-rank test: 2 D 148.000, df D 2, P < 0.0001; Wilcoxon test: 2 D 148.008, df D 2, P < 0.0001; Fig. 1A). Females that were treated with direct abamectin application, died within 24 h of treatment. This mortality rate was signiWcantly higher than that of the water treated females (Log-rank test: 2 D 115.000, df D 1, P < 0.0001; Wilcoxon test: 2 D 115.000, df D 1, P < 0.0001; ⬘ D 0.0166) or the untreated females (Log-rank test: 2 D 92.000, df D 1, P < 0.0001; Wilcoxon test: 2 D 92.000, df D 1, P < 0.0001; ⬘ D 0.0166). In contrast, there was no diVerence between water treated and untreated female survival (Log-rank test: 2 D 0.001, df D 1, P D 0.9743; Wilcoxon test: 2 D 0.029, df D 1, P D 0.8644; ⬘ D 0.0166). Similar to female survival, the abamectin treatment had a signiWcant negative eVect on male Diglyphus survival (Logrank test: 2 D 94.964, df D 2, P < 0.0001; Wilcoxon test: 2 D 93.731, df D 2, P < 0.0001; Fig. 1B). Males that were treated directly with abamectin died within 24 h of treatment. This mortality rate was signiWcantly higher than that of the water treated males (Log-rank test: 2 D 60.077, df D 1, P < 0.0001; Wilcoxon test: 2 D 60.077, df D 1, P < 0.0001; ⬘ D 0.0166) or the untreated males (Log-rank test: 2 D 63.000, df D 1, A
3.3. EVect of abamectin leaf residue on adult female Diglyphus When female Diglyphus were conWned within clipcages to chrysanthemum leaves that were treated with abamectin or water 24 h after application, abamectin had a signiWcant negative eVect on D. isaea adult female longevity (Log-rank test: 2 D 7.333, df D 1, P D 0.0068; Wilcoxon test: 2 D 12.971, df D 1, P D 0.0003; nabamectin D 22, nwater D 26; Fig. 3A). Although this was reduced Wve days after application, abamectin leaf residue still had a signiWcant eVect on female longevity (Log-rank test: 2 D 3.622, df D 1, P D 0.0570; Wilcoxon test: 2 D 3.569, df D 1, P D 0.0588; nabamectin D 24, nwater D 23; Fig. 3B). Longevity of females that were conWned to leaves seven (Log-rank test: 2 D 0.057, df D 1, P D 0.8117; Wilcoxon test: 2 D 0.007, df D 1, P D 0.9326;
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Fig. 3. EVect of leaf residue on adult female D. isaea longevity. (A) Residual toxicity eVect one day after application. (B) Residual toxicity eVect Wve days after application. (C) Residual toxicity eVect seven days after application. (D) Residual toxicity eVect 10 days after application.
nabamectin D 26, nwater D 29; Fig. 3C) and 10 days after application was not aVected by abamectin treatments (Log-rank test: 2 D 2.067, df D 1, P D 0.1505; Wilcoxon test: 2 D 0.276, df D 1, P D 0.5992; nabamectin D 25, nwater D 22; Fig. 3D).
3.4. EVect of abamectin application on Diglyphus larvae in the leaves Although Diglyphus emergence varied between blocks (F D 25.977, df D 1, P < 0.0001) under greenhouse condi-
100
1.0
90
0.9
80
0.8
70
0.7
60
177
0.6 Surviving
Mean Number of Diglyphus
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50 40 30
0.5 0.4 0.3
20
0.2
10
Water Abamectin
0.1
0
0.0
Water
Fig. 4. EVect of abamectin application on emergence of D. isaea adults from chrysanthemum plants in the greenhouse. Horizontal lines denote the mean (§SE) and diamonds 95% conWdence intervals.
tions, abamectin application did not signiWcantly aVect the numbers of emerged adult Diglyphus (F D 0.496, df D 1, P D 0.4834; Fig. 4). No interaction was detected between blocks and treatments (F D 2.052, df D 1, P D 0.1558). 3.5. EVect of abamectin applied to Diglyphus larvae in leaves on adult longevity When applied to leaves containing Diglyphus larvae, abamectin did not have a signiWcant eVect on the percentage of Diglyphus emerging (abamectin: 84.21% emerged; water: 92.59% emerged; P D 0.2692) or on the longevity of emerged Diglyphus adults (Log-rank test: 2 D 0.007, df D 1, P D 0.9309; Wilcoxon test: 2 D 0.1367, df D 1, P D 0.7116; Fig. 5). 3.6. EVect of direct abamectin contact with Diglyphus larvae We found that direct larval contact with abamectin had a signiWcant negative eVect on Diglyphus adult emergence (2 D 20.579, df D 2, P < 0.0001); 100% of abamectin treated Diglyphus did not survive (Fig. 6). Abamectin application signiWcantly aVected Diglyphus survival compared to the water treatment (P D 0.0004, ⬘ D 0.0166) and with the UTC treatment (P < 0.0001, ⬘ D 0.0166). No diVerence was found between the water and UTC treatments (P D 0.1237, ⬘ D 0.0166). 3.7. EVect of abamectin-poisoned leafminer larvae on Diglyphus larvae When untreated Diglyphus host larvae were replaced with larvae that were taken from plants treated with
1
2
3
4
5
6
Days Fig. 5. EVect of abamectin application on the longevity of D. isaea adults that emerged from the treated plants.
90 Percentage of adult emergence
Abamectin
A
80 70 60
A
50 40 30 20 10 0
B
Abamectin
Water
UTC
Fig. 6. EVect of direct abamectin contact with D. isaea larvae on the percentage of adult emergence. ‘UTC’ D untreated control. DiVerent letters indicate a signiWcant diVerence (P < 0.0166).
water or abamectin, abamectin-poisoned leafminer larvae had a signiWcant negative eVect on Diglyphus larvae survival (P < 0.0001); 91% of those larvae did not survive (Fig. 7).
4. Discussion Previous studies on the eVect of abamectin on Diglyphus spp. are confusing and conXicting. Abamectin was reported to have a “minimal eVect” on Diglyphus intermedius (Girault) and Diglyphus begini (Ashmead) in celery Welds (Trumble, 1985). Similarly, Hara (1986) found no abamectin adverse eVect on immature D. intermedius stages in cut chrysanthemum crops. In this study Hara (1986) found that D. isaea accounted for less than one percent of the total recovered parasitoid species, but no data were presented on D. isaea recov-
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Percentage of adult emergence
100 90 A
80 70 60 50 40 30 20 10
B
0 Abamectin
Water
Fig. 7. EVect of abamectin-poisoned leafminer larvae on D. isaea emergence percentage. DiVerent letters indicate a signiWcant diVerence (P < 0.05).
ery in the control vs. treated plots. In a celery Weld study, Weintraub (1999) showed that abamectin (plus 1% UltraWne oil) applications did not aVect the D. isaea adult population. In contrast, Schuster (1994) used laboratory tomato leaf-dip experiments and found that abamectin causes 73% D. intermedius larval mortality, and 38% D. intermedius pupa mortality. In a potato Weld study, Weintraub (2001) showed a reduction in the number of D. isaea immature stages only 14 days after a sole abamectin application. This D. isaea immature population started to recover 28 days from application and by the end of the study equaled the D. isaea immature population in the control plots. Prijono et al. (2004) and Schuster (1994) showed (in laboratory studies) that abamectin residue causes some mortality of D. isaea adults when they were exposed immediately after application. Our Wndings show that abamectin is lethal to the adult D. isaea (both sexes) when direct contact is made, meaning that an abamectin application in a greenhouse would eliminate most if not all D. isaea adults present. Moreover, adults that are released or emerge a few days later, and feed on food contaminated with abamectin (such as contaminated water or honeydew) would also be negatively aVected. Furthermore, parasitoid longevity would be negatively aVected if they contact recently (1–5 days) sprayed leaves. These results support previous studies that showed negative aVects of abamectin residue on Diglyhus (Schuster, 1994; Prijono et al., 2004). However, our results suggest that the abamectin residues were depleted 7 days after application under laboratory conditions and therefore a period of one week is suggested as the duration of time before releases of D. isaea into abamectin treated greenhouses can be made. Under Weld conditions, abamectin leaf residues can rapidly degrade (half-life < one day) probably caused by photodegradation (Bull et al., 1984). Greenhouses vastly vary in the extent to which
they block ultraviolet light. Therefore, abamectin residue depletion period may be shorter under some greenhouse conditions. These Wndings are similar to those of another leafminer parasitoid—Dacnusa sibirica (Shipp et al., 2000), where under cucumber greenhouse conditions adult mortality was less than 10% when they were exposed to abamectin treated leaves 6 days after application. Our results also show that abamectin is very toxic to D. isaea larvae when direct contact is made or when they feed on abamectin-poisoned leafminer larvae. Both of these experiments were done in highly manipulated laboratory experiments. These types of experiments were necessary to better understand the mechanism by which Diglyphus avoids abamectin poisoning. While they are inside the mine D. isaea larvae are protected and are not aVected by abamectin application (both under greenhouse and laboratory conditions). Abamectin probably does not penetrate the dead plant tissue and there is air surrounding both the parasitoid and its leafminer host. In addition, the hosts of immature D. isaea (i.e., leafminer larvae) are paralyzed or dead at this point, so they do not consume abamectin. Therefore, physical aspects of the mines and the fact that Diglyphus paralyzes its host provides an ecological refuge for D. isaea larvae from abamactin applications. Moreover, about 6–8 days after application (at 25 °C), adults should emerge into a relatively safe environment in terms of abamectin leaf residue. Abamectin application did not have a signiWcant eVect on D. isaea longevity after emerging from treated chrysanthemum plants. However, abamectin may cause some sub-lethal impacts on D. isaea (other than longevity) such as abnormal behavior, as well as decreased fecundity, health, and Wtness. These possible sub-lethal impacts of abamectin on D. isaea were not evaluated in this study. Despite being an excellent leafminer insecticide, our results suggest that abamectin is compatible with the parasitoid D. isaea for leafminer control in greenhouses provided that the duration of its residual toxicity is taken into account.
Acknowledgments We thank Thomas Costamagna, Bob Starnes, and Trinity Strehl for their invaluable advice and assistance to our study. We acknowledge the generous donation of Chrysanthemum cuttings by ‘Yoder Brothers,’ yellow sticky cards by ‘Seabright Laboratories,’ and Diglyphus by ‘Plant Sciences, Watsonville, CA, and ‘Koppert-Biological Systems.’ This research was supported by the National Research Initiative (NRI; USDA) Competitive Grants program, Proposal No. 2002-01971; and the American Floral Endowment.
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