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Poster 22: Mass-rearing Optimization of the Parasitoid Psyttalia lounsburyi for Biological Control of the Olive Fruit Fly F. Chardonnet1, A. Blanchet1, B. Hurtel1, F. Marini2, M.-C. Bon1, K.M. Daane3, C.H. Pickett4, X. Wang3 and L. Smith1 1
European Biological Control Laboratory, ARS-USDA, Campus international de Baillarguet, Montferrier-sur-Lez, Hérault, FRANCE,
[email protected] ;
[email protected] ;
[email protected] ;
[email protected] ,
[email protected], 2Biotechnology and Biological Control Agency, Roma, ITALY,
[email protected], 3University of California, Berkeley, California, USA,
[email protected],
[email protected], 4California Deptartment of Food and Agriculture, Meadowview, California, USA,
[email protected] The olive fruit fly, Bactrocera oleae (Rossi) (Diptera: Tephritidae), discovered in 1998 in California, is a direct pest of olives that has invaded the Mediterranean Region and California (Rice et al., 2003; Zalom et al., 2009). The fly is believed to have originated from Africa (Hoelmer et al., 2011), and Psyttalia lounsburyi Silvestri (Hymenoptera: Braconidae), a larval parasitoid from Africa, has been approved for release in the USA as a classical biological agent (Copeland et al., 2004; Daane et al., 2015). P. lounsburyi oviposits on B. oleae larvae inside the olive fruit, and completes development after the host pupates. However, it has been difficult to rear the parasitoid in the laboratory because it is multivoltine, and the host develops only in fresh olives, which are not available for most of the year (Daane et al., 2008). A method to rear the parasitoid on the factitious host, Mediterranean fruit fly, Ceratitis capitata (Wiedemannn) (Diptera: Teprhitidae), which can be reared on artificial diet throughout the year, was developed by Thaon et al. (2009). Although functional, this method needed to be improved to allow mass production indispensable for release in the field. Thus, we developed in our laboratory a number of ways to improve the efficiency of rearing by focussing on increasing productivity per female and reducing human labour. Thaon et al. (2009) developed an artificial 'olive' to stimulate the oviposition behaviour of P. lounsburyi female (Figs. P22.1 and P22.2). To create an “exposure ball”, a 2 cmdiameter polystyrene ball is completely covered with 2-mm thick layer of larval artificial diet. Seven-day-old larvae (third instar) of C. capitata are then placed all over the ball and maintain with a stretched sheet of Parafilm™. After a period of exposure, the “ball” device is removed from the parasitoid cage and larvae are placed in a larger aerated box with artificial diet until pupation. The parasitoids start emerging 21 and 23 days after parasitisation for males and females, respectively. To improve the use of this method and optimise production, we first determined the optimum duration of the period of exposure (3 hours instead of 24). We also estimated the best reproduction period for female parasitoid, which is between the first and the fifth week post-emergence. Thus, hosts are exposed 10 to ©CAB International 2017. Proceedings of the 5th International Symposium on Biological Control of Arthropods (eds. P.G. Mason, D.R. Gillespie and C. Vincent)
Poster 22: Mass-rearing Psyttalia lounsburyi – F. Chardonnet et al. (2017) 287
15 times to 1-2-week old mated females over a three-week period (instead of nine, i.e., during their whole life).
Fig. P22.1. Adult Psyttalia lounsburyi rearing cage. (photo: B. Hurtrel).
Fig. P22.2. A 5-mm diameter spherical shaped device for exposure of Ceratitis capitata larvae to Psyttalia lounsburyi. (photo: A. Blanchet).
The effect of high density (overcrowding) on reducing fecundity of entomophagous insects has long been known (Campos and Gonzales, 1991; Watt, 1960). For mass rearing, it is essential to be able to produce the maximum of descendants with the lowest number of breeder individuals. The previously applied method in our laboratory consisted of using a cohort of 200 parasitoid females and 100 males in a cylindric cage (25 cm diam., 20 cm length; i.e., 32.7 cm3 per wasp; Fig. P22.1). In order to reduce intra-specific competition, we reduced this density by putting 50 females and 25 males per cage (i.e. 130.8 cm3 per wasp). Thanks to this change, the number of descendants per female for one exposure increased by 48% (a mean of 2.11 descendants per female for 50-female cohorts vs. 1.42 for 200-female cohorts). In biological control programs using parasitoids, females are more valuable than males because males can fertilize multiple females, and only females directly kill the target pest. By reducing density, we were also able to improve the female sex ratio by 45% (increasing females from 33.1% to 47.9%). In order to standardize the P. lounsburyi rearing conditions, it was first essential to determine the number of fly larvae to expose to provide an ad libitum resource to parasitoids. Secondly, we had to find a simple and rapid method to collect and expose the chosen number of C. capitata larvae. For this study, we used samples of 7-day-old C. capitata larvae collected from “exposure balls” exposed to parasitoids. We measured the volume and the weight of each sample, and counted the number of individuals. We concluded that 1 ml of larvae contains about 662.4 individuals. Thus, for a cage containing 50 P. lounsburyi females with one exposure ball covered with 2 ml of larvae, there are about 26 host available per female. The maximum number of descendants per female during a 3 hours oviposition period was 5. So, 2 ml of larvae easily provided an "ad libitum" condition for parasitoid oviposition. To standardize the use of a chosen amount of fly larvae for exposure, we created a milliliter "dip net" made of a Falcon milliliter tube (15 ml tube with conic bottom). The bottom of the tube was cut and a thin nylon mesh was fixed with hot glue on one end, which permits scooping larvae from water (the simplest method of handling) while
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Poster 22: Mass-rearing Psyttalia lounsburyi – F. Chardonnet et al. (2017)
allowing the liquid to pass through. A plastic handle was also fixed to the device for easier handling (Fig. P22.3).
Fig. P22.3. Milliliter "dip net" to measure 2ml of fly larvae. (photo: F. Chardonnet).
Counting and sexing P. lounsburyi adults in preparation for shipment was one of the most time-consuming tasks during mass production (limiting the increase of production). In order to reduce this workload, we tested the reliability of using subsamples of about 10% of the fly pupae produced (after exposure to parasitism), from which the total number of emerged parasitoid adults is estimated. We counted the total number of emerged individuals and compared this number with the estimation made from the 10% sample. Because the estimate did not significantly differ from the real number of parasitoids collected, we considered that using a subsample of 10% of pupae was an accurate method to estimate adult production. This reduced counting time by 90 %, saving time which could be used to further increase production. Thanks to all these modifications, we significantly improved the number of progeny produce per female, and the female sex ratio of progeny. We also improved conditions for rearing larvae and holding emerged adults until release and thus improved survivorship of both immatures and adults. In 2016, we were able to produce over 54,000 adults, and shipped over 22,900 for release in California. References Campos, M. and Gonzalez, R. (1991) Effect of parent density on fecundity of two parasitoids (Hymenoptera: Pteromalidae) of the olive beetle, Phloeotribus scarabaeoides (Coleoptera: Scolytidae). Entomophaga, 36, 473-480. Copeland, R.S., White, I.M., Okumu, M., Machera, P. and Wharton, R.A. (2004) Insects associated with fruits of the Oleaceae (Asteridae, Lamiales) in Kenya, with special reference to the Tephritidae (Diptera). Bishop Museum Bulletins in Entomology, 12, 135-164. Daane, K.M., Sime, K.R., Wang, X.-G., Nadel, H., Johnson, M.W. and Walton, V.M. (2008) Psyttalia lounsburyi (Hymenoptera: Braconidae), potential biological control agent for the olive fruit fly in California. Biological Control, 44, 78–89. Daane, K.M., Wang, X., Nieto, D.J., Pickett, C.H., Hoelmer, K.A., Blanchet, A. and Johnson, M.W. (2015) Classic biological control of olive fruit fly in California, USA: release and recovery of introduced parasitoids. BioControl, 60, 317-330.
Poster 22: Mass-rearing Psyttalia lounsburyi – F. Chardonnet et al. (2017) 289 Hoelmer, K.A., Kirk, A.A., Pickett, C.H., Daane, K.M. and Johnson, M.W. (2011) Prospects for improving biological control of olive fruit fly, Bactrocera oleae (Diptera: Tephritidae), with introduced parasitoids (Hymenoptera). Biocontrol Science and Technology, 21, 1005-1025. Rice, R., Phillips, P., Stewart-Leslie, J. and Sibbett, G. (2003) Olive fruit fly populations measured in central and southern California. California Agriculture, 57, 122–127 Thaon, M.., Arnaud, B. and Nicolas R. (2009) Contribution à l’optimisation de l’élevage du parasitoïde Psyttalia lounsburyi. Cahier des Techniques Inra, 66, 21-31. Watt, K. E. F. (1960) The Effect of Population Density on Fecundity in Insects. The Canadian Entomologist, 92, 674-695. Zalom, F.G., Van Steenwyk, R.A., Burrack, H.J. and Johnson, M.W. (2009) Pest notes: olive fruit fly. University of California Agriculture and Natural Resources, Publication 7411.
