The authors thank Lukas Seehausen and Marc Kenis (CABI, Switzerland) for kindly providing G1 G. brasiliensis. Funding in Italy was provided by Provincia Autonoma di Trento, Trento, Italy, and in the US by the National Institute of Food and Agriculture, USDA Specialty Crops Research Initiative award (#2020-5118-32140), USDA Animal and Plant Health Inspection Service (Farm Bill, fund 14-8130-0463), and USDA ARS CRIS base funds (project 8010-22000-033-00D). The USDA is an equal-opportunity provider and employer and does not endorse products mentioned in this publication.

1. Methods for small-scale laboratory rearing of G3 Ganaspis brasiliensis
<ol>
	<li>Prepare host diet.
	<ol>
		<li>Add 600 mL of distilled water to a 1,500 mL glass container, and heat the water on a hot plate.</li>
		<li>Add 88.6 g of commercially available dry diet (made of agar, Brewer&#39;s yeast, corn meal, methyl paraben, and sucrose) or prepare diet using the formula published by Dalton et al.28 (see step 2.1.2).</li>
		<li>Add 300 mL of distilled water into the dry diet and stir the diet mixture thoroughly.</li>
		<li>Add the mixture to the boiling water.</li>
		<li>Allow the liquid diet on the hot plate to boil for 10 min while periodically stirring the mixture to prevent it from burning.</li>
		<li>Allow the diet to cool at room temperature for 30 min while stirring it occasionally to distribute the release of heat evenly and prevent the diet from solidifying on the surface.</li>
		<li>Measure 6.7 mL of 95% EtOH in one container and 3.5 mL of 1 M propionic acid solution in another container.</li>
		<li>Once the diet has cooled, add the EtOH and then the propionic acid solution, stirring thoroughly after each addition.</li>
		<li>Prepare blueberries (purchased from the local market) by rinsing them in cold water, then in a sodium hypochlorite bleach solution (diluted to 5%), and cold water again.</li>
		<li>Dry the fruit with a paper towel and manually mash them until the skin on each fruit is broken and the juices and flesh of the fruit are exposed.</li>
		<li>Add 25-30 g of mashed blueberries to each 250 mL flask. Tap the sides of the flask to ensure the interior bottom of the flask is covered with an even layer of mashed blueberries.</li>
		<li>Pour the prepared diet into each flask so that it just covers the top of the mashed blueberries.</li>
		<li>Add foam stoppers to the necks of the flasks and allow the diet to solidify at room temperature (Figure 1).</li>
		<li>Once the diet has solidified, use it immediately or store at 5 &#176;C for up to 3 weeks.</li>
	</ol>
	</li>
	<li>Rear host Drosophila suzukii.
	<ol>
		<li>Remove the stored diet from the refrigerator and allow it to equilibrate to the ambient room temperature, or use freshly made diet.</li>
		<li>Cut a piece of absorbent paper towel (e.g., 5 cm x 20 cm) and twist it in the center. Place the twisted middle section of the paper towel in the flask (Figure 1).</li>
		<li>Wet the paper towel and surface of the diet with distilled water to retain moisture.</li>
		<li>Transfer sexually mature adult flies from the current colony fly flasks to a new diet flask by carefully removing the stopper on the old flask and quickly inverting the flask and aligning the opening of the old flask with the new flask.</li>
		<li>Gently tap on the side of the old flask to induce the flies to drop into the new flask. Ensure that there are ~25-30 mating pairs of D. suzukii in the new flask. Once there are enough flies in the new flask, quickly flip the old flask upright and replace the stoppers on both flasks.</li>
		<li>Repeat the transfers of flies until no flies are left in the old flasks. If necessary, combine or collect flies from more than one old flask into a new flask to ensure there are enough flies (20-30 pairs) per flask.</li>
		<li>Hold the new flasks after a week of exposure to adult flies at suitable conditions (21 &#176;C, 16 L: 8 D photoperiod, 60%-80% relative humidity [RH%]) in an environmental chamber for 3 weeks for fly emergence.</li>
	</ol>
	</li>
	<li>Expose host larvae to parasitoids.
	<ol>
		<li>Take a flask (see step 1.2.7) containing fly eggs and larvae after removing any adult flies and the twisted paper towel from the flask.</li>
		<li>Fold a piece of absorbent paper towel in half and put it in the flask as a pupation substrate for parasitized larvae.</li>
		<li>Aspirate six female and male pairs of G3 G. brasiliensis into each flask (Figure 1). Streak a thin layer of honey on the bottom of the foam stopper.</li>
		<li>Leave the parasitoids in the flask for 5 days.</li>
		<li>After a 5-day exposure, remove the parasitoids and hold the flasks under conditions described above in an environmental chamber for 35 days until the expected wasp emergence.</li>
	</ol>
	</li>
	<li>Collect and store adult parasitoids.
	<ol>
		<li>During the second and third weeks of incubation, check the flasks weekly for early host emergence and remove the adult flies.</li>
		<li>Once adult parasitoids start to emerge, aspirate them three times per week and hold them in drosophila vials (e.g., 2.5 cm x 9.5 cm) (Figure 1).</li>
		<li>Place a small piece of paper towel moistened, but not saturated, with distilled water at the bottom of the vial.</li>
		<li>Add ~60 parasitoids to each vial and label the vial with the emergence dates. Streak a thin layer of honey on the bottom of the foam stopper, twice per week. Store the vials with adult parasitoids under the conditions described above in the environmental chamber for up to a month if not used sooner.</li>
		<li>Remoisten the paper in the vial once every 4-7 days or replace the paper towel if there are signs of mold.</li>
	</ol>
	</li>
</ol>
2. Methods for large-scale rearing of G1 Ganaspis brasiliensis
<ol>
	<li>Implement a large-scale rearing of host Drosophila suzukii.
	<ol>
		<li>Rear D. suzukii within large, knitted mesh-covered cages (e.g., 50 cm x 50 cm x 100 cm) each containing 1,500-2,000 sexually mature adult flies (sex ratio 50:50) (Figure 2).</li>
		<li>Prepare the Standard Drosophila Medium (SDM) by boiling all the ingredients (6 g of bacteriological agar, 75 g of cornmeal, 17 g of nutritional yeast, 15 g of saccharose, 10 g of soybean flour, 10 mL of propionic acid) in 1 L of distilled water for 10 min while periodically stirring the mixture to prevent it from burning28.</li>
		<li>Let the mixture cool down for 5 min and add 5 g of ascorbic acid.</li>
		<li>Pour the freshly cooked SDM into 9 cm Petri dishes and allow the medium to solidify at room temperature before closing the plates.</li>
		<li>Stack up the SDM Petri dishes, wrap the stack with aluminum foil, and store the dishes at 4 &#176;C for up to 2 weeks.</li>
		<li>Within each rearing cage, place a plate with water-soaked cotton and four to six Petri dishes with SDM (Figure 2).</li>
		<li>Twice per week, replace the infested SDM Petri dishes with fresh ones.</li>
		<li>Place the infested SDM Petri dishes without lids individually into plastic cups (13.3 cm diameter or 800 mL), close each cup with a covering of fine mesh (&#60;0.5 mm), and incubate for 12-15 days at 23 &#176;C and 75% RH (Figure 2).</li>
		<li>Transfer the newly hatched D. suzukii adults from the plastic cups to the rearing cages.</li>
	</ol>
	</li>
	<li>Prepare host larvae.
	<ol>
		<li>Rinse the blueberries in cold water for 1 min, and soak the fruits in a basin filled with a bleach solution (diluted to 5%) for 3 min.</li>
		<li>Drain the bleach solution and fill the basin with cold water to rinse the blueberries. Mix gently by hand for at least 30 s.</li>
		<li>Repeat step 2.2.2 with fresh water at least three times to remove bleach residues and other arthropods (e.g., mites, thrips) that may be present on the fruit.</li>
		<li>Place the fruit on a tray with several layers of absorbent paper towels and carefully tilt the tray back and forth, rolling the berries around to dry them.</li>
		<li>Prepare several 9 cm Petri dishes (either the top or the bottom halves, facing up) and fill each one with the washed blueberries (15-25 fruits per plate depending on the fruit size).</li>
		<li>During late afternoon hours, expose the Petri dishes to sexually mature adult flies within the host rearing cages (see step 2.1) and leave them overnight.</li>
		<li>The next morning, remove the Petri dishes from the host rearing cages by gently blowing or tapping on them to dislodge the flies on the fruits and use the infested fruit for the rearing of parasitoids (see step 2.4).</li>
	</ol>
	</li>
	<li>Implement a large-scale parasitoid rearing.
	<ol>
		<li>Use two types of cages to rear the parasitoid: one for parasitism and another for wasp emergence.</li>
		<li>Ensure that the parasitism cage is cubic (e.g., 45 cm each side) with a clear plastic panel on the front for observing insect activity, two 18 cm sleeve openings in the front panel for the addition or removal of insects and the replacement of food material, and fine polyester netting (e.g., 96 x 26 mesh) on the top and sides for ventilation.</li>
		<li>Make the emergence cage smaller (e.g., 30 cm each side), with a single sleeve opening on two opposite sides and a clear plastic panel on the front for visibility (Figure 2).</li>
		<li>Ensure that both cage types have a thin string that hangs below the ceiling from which to suspend one to several feeders (Figure 2). 
		NOTE: A feeder consists of a large cylindric foam stopper (9 cm diameter) covered with scattered honey droplets, and can be placed on the cage floor or hung from the cage ceiling (Figure 2).</li>
		<li>Within each cage, provide water in a straight-wall drosophila vial (2.5 cm x 9.5 cm) sealed with a cellulose acetate plug (2.5 cm diameter) every 5-7 days depending on the RH. Hang the vial upside-down from the cage ceiling (Figure 2).</li>
	</ol>
	</li>
	<li>Expose the host larvae to the parasitoids.
	<ol>
		<li>Expose the host-infested fruit within the Petri dishes to G1 G. brasiliensis immediately after D. suzukii overnight infestation (see step 2.2.7).</li>
		<li>Leave the 10-15 Petri dishes of infested fruit in the parasitization cage containing 1,500-2,000 wasps for 2-3 days.</li>
		<li>Use plastic cups (13.3 cm diameter or 800 mL) with layers of absorbent paper on the bottom to collect the fruit containing the parasitized hosts (Figure 2).</li>
		<li>Place the open cups in the eclosion cage and incubate for at least 28 days at 21 &#176;C and 65% RH (Figure 2).</li>
		<li>During the second and third weeks of incubation, check the cage weekly for early host eclosion and remove the adult flies to facilitate the successive collection of parasitoids.</li>
		<li>At the end of the fourth week of incubation, add a feeder and a water source to the cage.</li>
	</ol>
	</li>
	<li>Collect and store the adult parasitoids.
	<ol>
		<li>Once parasitoid emergence starts, collect a portion (10%-15%) of the adults and transfer them back to the parasitism cage to replace old unproductive individuals.</li>
		<li>Collect and store the remaining parasitoids in plastic cups (13.3 cm diameter or 800 mL) (Figure 3A).</li>
		<li>Place a tube (2 mL) filled with water and sealed with a dental cotton roll (1 cm x 3.8 cm) at the bottom of the cup (Figure 3A).</li>
		<li>Close the cup with a modified lid fitted with a removable foam stopper (3.5 cm diameter) as a feeder substrate and a mesh-covered hole for ventilation (Figure 3B).</li>
		<li>Add up to 700 adults to each cup (sex ratio 50:50), label the cup with the emergence date, and store it in an environmental chamber (17 &#176;C; 65% RH) until used, or for up to 1 month (Figure 3B).</li>
	</ol>
	</li>
	<li>Ship the adult parasitoids.
	<ol>
		<li>Use conical tubes (50 mL) to ship the adult parasitoids.</li>
		<li>Pierce a ventilation hole (8 mm diameter) on the cap and cover it with a fine mesh net (Figure 3C).</li>
		<li>Add a cellulose acetate feeding ring on the inside of the cap (Figure 3C).</li>
		<li>Prepare a saturated sucrose solution using distilled water, apply a few drops on the feeding ring, and let it absorb the liquid.</li>
		<li>Place a fan-shaped piece of absorbent paper towel within the tube (Figure 3D).</li>
		<li>Add ~200 adult parasitoids to each tube, and place the tubes in an insulated shipping container together with ice packs.</li>
	</ol>
	</li>
</ol>

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T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laboratory+survival+of+Drosophila+suzukii+under+simulated+winter+conditions+of+the+Pacific+Northwest+and+seasonal+field+trapping+in+five+primary+regions+of+small+and+stone+fruit+production+in+the+United+States.">Laboratory survival of Drosophila suzukii under simulated winter conditions of the Pacific Northwest and seasonal field trapping in five primary regions of small and stone fruit production in the United States.</a> Pest Managagement Science. 67 (11), 1368-1374 (2011).</li></ol>

The authors have no conflicts of interest to disclose.

Long-term research and subsequent field releases of a biological control agent depend on the availability of effective and economical rearing techniques. The described methods in this study have proven to be efficient protocols for both small-scale and large-scale rearing of Ganaspis brasiliensis. The small-scale rearing protocol has been developed over several years to optimize the amount of labor and reduce specialized equipment needed to maintain the parasitoid and host colonies simultaneously. It is suitable for maintaining a colony for laboratory research or bioassays. Similar methods have been used by the authors to rear this parasitoid for quarantine evaluations of this parasitoid. The large-scale rearing protocol will allow to produce large numbers of wasps for field release, as were recently conducted in Italy. These technologies can easily be transferred to other laboratories, producers, or companies for large-scale rearing in the near future, as well as serving as the basis for further improvements in methodologies.
These protocols may also be used to rear Leptopilina japonica, as both Ganaspis brasiliensis and L. japonica are similar in terms of their host habitat preference7,8, host stage preference or reproductive biology15, thermal performance18, and foraging efficiency in laboratory bioassays13,15, as well as behavioral responses towards host-associated cues12, except that L. japonica seems to have a broader host range than that of even G3 G. brasiliensis20. Both parasitoids have been reared using similar methods as described herein. For small-scale rearing, both parasitoids can also be reared in drosophila vials, typically by transferring 20 young (1-2 days old) Drosophila suzukii larvae to a drosophila vial filled with 2 cm of artificial diet, or placing two infested fruits, each containing 5-10 young D. suzukii larvae, and exposing them to two mated female wasps for 2-3 days, both producing ~10 offspring per vial13,15,22.
As discussed above, the G1 and G3 Ganaspis brasiliensis used in this protocol may differ slightly in some host-searching behaviors and host specificity21,22. Girod et al.21 report that the Japanese G1 G. brasiliensis was more strictly specific to Drosophila suzukii,&#160;and did not appear to perform well in pure artificial diet in vials compared to its performance on host fruit. Matsuura et al.25 also reported that G1 G. brasiliensis populations collected from D. suzukii-infested cherries in Japan specialize on D. suzukii. The small-scale rearing method using diet mixed with blueberries was developed initially for rearing G3 G. brasiliensis because G1 G. brasiliensis was not available at that time. Later, this method was found to not work well for the rearing of G1 G. brasiliensis (Wang et al. unpublished data).
Therefore, for small-scale rearing of G1 G. brasiliensis, it is suggested to modify the host substrate by (1) exposing the flies directly to blueberries (or other host fruit) to collect host larvae in the fruit, and (2) transferring exposed fruit to a standard Drosophila culture medium for the larvae to develop in a low-competition environment by placing the infested fruits containing the host larvae onto the diet in the flasks for exposure to the parasitoids. This will allow the parasitized larvae to feed on the diet, especially at high host densities, and the parasitized host pupae to be collected from the paper towel. Alternatively, G1 G. brasiliensis can be reared on fruit directly in plastic containers (various sizes) by exposing 5-10 female wasps to 10-20 infested blueberries for 4-5 days, which produced up to 80 offspring per container, depending on host density (Wang et al., unpublished data). For this method, it is recommended to place the infested fruit on a raised metal mesh (&#34;hardware cloth&#34;) to allow the parasitoids to access the infested fruit from all directions, especially if too many fruits are placed in one container. Ideally, one or two layers of infested fruit should be placed in each container for this rearing method. These alternative small-scale methods should also work well for the rearing of G3 G. brasiliensis.
Regardless of the rearing methods and scales (vial, flask, container, or cage), it is critical to maintain suitable temperature, humidity, as well as control host or female wasp age, density, and exposure time for the rearing of both G1 Ganaspis brasiliensis and G3 G. brasiliensis. Drosophila suzukii larvae developed in about 1 week under normal laboratory conditions (e.g., 22 &#177; 2 &#176;C) 15. Female G. brasiliensis preferred to attack younger (1-2 day old) than older (3-4 day old) host larvae, although various ages of host larvae could be attacked15. At 22 &#177; 2 &#176;C, G. brasiliensis females emerged with a substantially high proportion (~50%) of their lifetime complement of mature eggs, and the mature egg load reached a peak after 2-3 days15. Adult females survived ~20 days when provided unlimited access to hosts and began oviposition within 2 days after emergence, reaching a peak of oviposition within 5-10 days and thereafter gradually reducing oviposition15. Therefore, young (&#60;10 days old) female wasps should be used in the rearing, but female wasps could be reused for the rearing when they are in a short supply. The parasitoid could readily develop at 21-25 &#176;C, but temperatures below 17.2 &#176;C appeared to trigger a facultative diapause17. It is therefore recommended to use a temperature range of 21-25 &#176;C for optimal development of both the fly and the parasitoid.
Further, exposure time of more than 5 days will not likely increase the productivity of the parasitoid. Increased parasitoid density based on the small-scale rearing for G3 G. brasiliensis appears to not increase the productivity, possibly due to mutual interference among foraging female wasps. Six male-female pairs and a 5-day exposure time seem to be an ideal combination for the laboratory small-scale rearing, although the rearing methods may be improved in the future by further optimizing the ratio of host to parasitoid. A major mortality factor for the parasitoid seems to be related to low humidity, as many parasitoids were observed to be unable to eclose with dry substrate conditions. Adding a piece of absorbent paper towel underneath the fruit not only absorbs juices as the fruit degrades, but also provides a substrate that can be dampened to increase humidity or provide a pupation substrate for the fly.
For the small-scale rearing, a flask can maintain humidity better than a vial because the former has a narrow neck. It was also found in this study that fresh blueberries coated with a dusting of active dry yeast helped prevent the formation of mold and enhanced the attraction of the fruit to the flies. Other aspects of the parasitoid&#160;rearing that remain to be explored include (1) the possibility of rearing this parasitoid on alternative hosts or host fruits and how alternative hosts or host fruits would affect the parasitoid&#39;s efficiency, (2) factors affecting the parasitoid&#39;s offspring fitness and sex ratio, (3) the ability of this parasitoid (both G1 and G3 ) to adapt to artificial diet conditions, and (4) the genetic or behavioral changes that might occur with adaptation.

Native to East Asia, the spotted-wing drosophila, Drosophila suzukii (Matsumura) (Diptera: Drosophilidae), has established widely in the Americas, Europe, and parts of Africa1,2. The fly is extremely polyphagous, being capable of utilizing various cultivated and wild fruits with soft and thin skins in its native and invaded regions1,2,3. Current management strategies for this pest rely heavily on the frequent use of insecticides that target adult flies in crop fields when susceptible fruit are ripening. Repeated sprays are often used, possibly due to consistent spillover of reservoir fly populations from non-crop habitats and lack of effective natural enemies resident in the invaded regions1,4. Biological control, especially by means of self-perpetuating specialized parasitoids, may help suppress fly populations at the landscape level and play a critical role for sustainable area-wide management of this highly mobile and polyphagous pest4,5,6.
Over the past decade, researchers have focused efforts to discover co-evolved parasitoids of Drosophila suzukii in the fly&#39;s native ranges in East Asia7,8,9, as well as effective but newly associated parasitoids in the fly&#39;s invaded regions in the Americas and Europe4,5,6. In the fly&#39;s newly invaded regions, commonly occurring larval Drosophila parasitoids, such as Asobara c.f. tabida (Nees) (Hymenoptera: Braconidae), Leptopilina boulardi (Barbotin et al.), and L. heterotoma (Thompson) (Hymenoptera: Figitidae), are unable to develop from or have low parasitism levels on D. suzukii due to the fly&#39;s strong immune resistance10. Only some cosmopolitan and generalist pupal parasitoids such as Pachycrepoideus vindemiae (Rondani) (Hymenoptera: Pteromalidae) and Trichopria drosophilae (Perkins) (Hymenoptera: Diapriidae) in North America and Europe, and Trichopria anastrephae Lima in South America can readily develop from this fly4. In contrast, explorations in East Asia have discovered a number of larval parasitoids from D. suzukii4,5,6. Among them, Asobara japonica Belokobylskij, Ganaspis brasiliensis Ihering, and Leptopilina japonica Novkovi&#263; &#38; Kimura are the dominant larval parasitoids7,8,9,11. In particular, the two figitids (L. japonica and G. brasiliensis) were the major parasitoids predominantly found in fresh fruits infested by D. suzukii and/or other closely related drosophilids in natural vegetation7,8,9. These three Asian larval parasitoids were imported to quarantine facilities in the USA and Europe, and evaluated for their relative efficiency12,13,14,15,16,17, climatic adaptability18, potential interspecific competitive interactions19, and, most importantly, host specificity8,20,21,22.
Quarantine evaluations showed that Ganaspis brasiliensis was more host-specific to Drosophila suzukii than other tested Asian larval parasitoids, although it likely consists of different biotypes or cryptic species with varying host specificity8,21,22,23,24. Nomano et al.22 grouped Ganaspis individuals from different geographical regions into five genetic groups (named as G1-G5) based on molecular analyses of the mitochondrial cytochrome oxidase I gene fragment. The G2 and G4 groups are reported only from a few south Asian tropical locations, and the G5 group was reported from Asia and other regions (e.g., Argentina, Brazil, Hawaii, and Mexico) from unknown host(s) (Buffington, personal observation). Field collections of wild fruits infested by D. suzukii in South Korea7, China8, and Japan9,23,25 found G1 alone or a mixture of specimens representing groups G1 and G3. The two groups seem to be sympatric and co-exist on the same host plants inhabited by D. suzukii and other closely related host flies. Nonetheless, some differences have been observed between the two groups, with G1 seemingly having a higher degree of host- or host-habitat-specificity to D. suzukii than G3, although they both attack a number of closely related species in the quarantine tests21,22. Further detailed molecular analyses may help determine the species status, especially for the G1 and G3 groups. This study refers to them as G1 G. brasiliensis and G3 G. brasiliensis. Some early studies also named the G1 G. brasiliensis as G. cf. brasiliensis14,21,22. The G1 G. brasiliensis has recently been approved for field release against D. suzukii in the USA and Italy (several other European countries are also currently considering its introduction), while the G3 G. brasiliensis may be considered for field release in the near future. Recent surveys also found adventive populations of both L. japonica and G1 G. brasiliensis in British Columbia, Canada26, and Washington State, USA (Beers et al., unpublished data), and adventive L. japonica populations in Trento province, Italy27.
Given the significant interest in the development of biological control programs for Drosophila suzukii management and the substantial biological control potential of adventive and deliberate introductions of Ganaspis brasiliensis, there is a need to develop efficient rearing methods for this larval parasitoid for future long-term research and/or field release. This protocol and associated video article describe two sets of rearing methods for this parasitoid: (1) small-scale laboratory rearing in flasks using a mixture of host fruit (blueberry) and artificial diet for the culture of D. suzukii. The methods were developed using G3 material originally collected from Kunming, China8. (2) Mass rearing for field release in large cages using host fruit (blueberry) for the culture of D. suzukii. The genetic group used for the large-scale rearing was G1 stock originating in Tokyo, Japan9,22. Other scales of rearing methods, such as using vials or small containers for both groups, are also briefly discussed.

<table><tbody><tr><td>Active dry yeast</td><td>Fleischmanns Yeast, Cincinatti, OH, USA</td><td>None</td><td>Used to cover fruit to reduce mold growth and enhance the frui attraction to the flies</td></tr><tr><td>Bacteriological agar</td><td>Merk Life Science S.r.l., Milan, Italy</td><td>A1296 - 5KG</td><td>Used to prepare the Standard Drosophila Medium</td></tr><tr><td>Bleach solution</td><td>Clorox Company, Oakland, CA, USA</td><td>None</td><td>Used to disinfect flesh fruit</td></tr><tr><td>Blue stopper</td><td>Azer Scientific, Morgantown, PA, USA</td><td>ES3837</td><td>Used for sealing the tube while allowing ventilation for insects</td></tr><tr><td>Blueberries</td><td>Grocery Store, Newark, DE, USA</td><td>None</td><td>Provided as host fruit for the flies (various other fruit can also be used)</td></tr><tr><td>BugDorm insect rearing cage (W24.5 x D24.5 x H63.0 cm)</td><td>Mega View Science Co. Ltd., Taichung, Taiwan</td><td>4E3030</td><td>Used for rearing parasitoids (parasitism cage)</td></tr><tr><td>BugDorm insect rearing cage (W32.5 x D32.5 x H32.5 cm)</td><td>Mega View Science Co. Ltd., Taichung, Taiwan</td><td>4E4590</td><td>Used for rearing flies</td></tr><tr><td>BugDorm insect rearing cage (W32.5 x D32.5 x H32.5 cm)</td><td>Mega View Science Co. Ltd., Taichung, Taiwan</td><td>4E4545</td><td>Used for rearing parasitoids (eclosion cage)</td></tr><tr><td>Chicken wire (0.64 cm, 19 gauge)</td><td>Everbilt, OH, USA</td><td>308231EB</td><td>Used to lift up the fruit to allow maximum parasitoid oviposition</td></tr><tr><td>Cornmeal</td><td>Grocery Store, Trento, TN, Italy</td><td>None</td><td>Used to prepare the Standard Drosophila Medium</td></tr><tr><td>Dental cotton roll (1 x 3.8 cm)</td><td>Gima S.p.A., Gessate, MI, Italy</td><td>35000</td><td>Used for providing water to the parasitoids within the storage container</td></tr><tr><td>Drosophila diet</td><td>Frontier Scientific, Newark, DE, USA</td><td>TF1003</td><td>Custom diet used to rear flies</td></tr><tr><td>Drosophila vial narrow, Polystirene (2.5 x 9.5 cm)</td><td>VWR International, LLC., Radnor, PA, US</td><td>75813-160</td><td>Used for providing water to the parasitoids within the cage</td></tr><tr><td>Drosophila vial plugs, Cellulose acetate (2.5 cm)</td><td>VWR International, LLC., Radnor, PA, US</td><td>89168-886</td><td>Used for providing water to the parasitoids within the cage</td></tr><tr><td>Erlenmeyer flask (250 mL)</td><td>Carolina Biological, Burlington, NC, USA</td><td>731029</td><td>Used for rearing flies and parasitoids</td></tr><tr><td>Falcon-style centrifuge tube (50 mL)</td><td>VWR International, LLC., Radnor, PA, US</td><td>VWRI525-0611</td><td>Modified to ship adult parasitoids</td></tr><tr><td>Foam stopper</td><td>Jaece Industries, North Tanawanda, NY, USA</td><td>L800-C</td><td>Used for sealing the flasks while allowing ventilation for insects</td></tr><tr><td>Honey</td><td>Grocery Store, Newark, DE, USA</td><td>None</td><td>Provided as food for parasitoids</td></tr><tr><td>Identi-Plug plastic foam stopper</td><td>Fisher Scientific Company, L.L.C., Pittsburg, PA, US</td><td>14-127-40E</td><td>Used as feeder for parasitoids and to seal the storage container</td></tr><tr><td>Industrial paper towel</td><td>Grocery Store, Newark, DE, USA</td><td>None</td><td>Provided as a pupation substrate for pupae and mitigated moisture</td></tr><tr><td>Micron mesh fabric (250 mL)</td><td>Industrial Netting, Maple Grove, MN, USA</td><td>WN0250-72</td><td>Used to make ventilation lid for insects</td></tr><tr><td>Nutritional yeast (flakes)</td><td>Grocery Store, Trento, TN, Italy</td><td>None</td><td>Used to prepare the Standard Drosophila Medium</td></tr><tr><td>Paper coaster (10.2 cm)</td><td>Hoffmaster, WI, USA</td><td>35NG26</td><td>Porvided as pupation substrate for flies and parsitized pupae</td></tr><tr><td>Plastic cup (&amp;Oslash; 13.3 cm, 800 mL)</td><td>Berry Superfos, Taastrup, Denmark</td><td>Unipak 5134</td><td>Modified to store adult parasitoids</td></tr><tr><td>Plastic lid (&amp;Oslash; 13.3 cm)</td><td>Berry Superfos, Taastrup, Denmark</td><td>PP 2830</td><td>Modified to store adult parasitoids</td></tr><tr><td>Propionic acid</td><td>Merk Life Science S.r.l., Milan, Italy</td><td>P1386 - 1L</td><td>Used to prepare the Standard Drosophila Medium</td></tr><tr><td>Saccharose</td><td>Grocery Store, Trento, TN, Italy</td><td>None</td><td>Used to prepare the Standard Drosophila Medium</td></tr><tr><td>Soup cup with lid (475 mL)</td><td>StackMan, Vietnam</td><td>DC1648</td><td>Used for parasitized larvae to pupate</td></tr><tr><td>Soybean flour</td><td>Grocery Store, Trento, TN, Italy</td><td>None</td><td>Used to prepare the Standard Drosophila Medium</td></tr><tr><td>White felt washer (0.64 cm thick, 5 mm ID x 20 mm OD)</td><td>Quiklok, Lincoln, NH, US</td><td>WFW/.25 x 5 x 20 mm</td><td>Used as feeding ring for parasitoids</td></tr></tbody></table>

methods for rearing the parasitoid ganaspis brasiliensis, a promising biological control agent for the invasive drosophila suzukii

Native to East Asia, the spotted-wing drosophila, Drosophila suzukii (Matsumura) (Diptera: Drosophilidae), has established widely in the Americas, Europe, and parts of Africa over the last decade, becoming a devastating pest of various soft-skinned fruits in its invaded regions. Biological control, especially by means of self-perpetuating and specialized parasitoids, is expected to be a viable option for sustainable area-wide management of this highly mobile and polyphagous pest. Ganaspis brasiliensis Ihering (Hymenoptera: Figitidae) is a larval parasitoid that is widely distributed in East Asia, and has been found to be one of the most effective parasitoids of D. suzukii.
Following rigorous pre-introduction evaluations of its efficacy and potential non-target risks, one of the more host-specific genetic groups of this species (G1 G. brasiliensis) has been approved recently for introduction and field release in the United States and Italy. Another genetic group (G3 G. brasiliensis), which was also commonly found to attack D. suzukii in East Asia, may be considered for introduction in the near future. There is currently enormous interest in rearing G. brasiliensis for research or in mass-production for field release against D. suzukii. This protocol and associated video article describe effective rearing methods for this parasitoid, both on a small scale for research and a large scale for mass-production and field release. These methods may benefit further long-term research and use of this Asian-native parasitoid as a promising biological control agent for this global invasive pest.

Figure 4 shows representative results of the small-scale laboratory rearing of G3 Ganaspis brasiliensis using two different parasitoid densities (six or 10 pairs) and two different exposure times (5 or 10 days) at the quarantine facility of the USDA-ARS Beneficial Insects Introduction Unit (Newark, Delaware). There were 14 replicates for each combination of parasitoid density and exposure time. In total, the 64 flasks produced 4,018 wasps (71.7 &#177; 4.9 offspring per flask) with 49.5% &#177; 1.9% female offspring. At 21 &#176;C, adult parasitoids emerged approximately 30-35 days after oviposition. The parasitoid density and exposure time did not significantly affect the total number of offspring produced per replicate (flask) (one-way ANOVA, F3,52 = 0.379, P = 0.769) and had only a marginal effect on the percentage of female offspring (one-way ANOVA, data were logit-transformed prior to the analysis as needed to stabilize the variation, F3,52 = 2.796, P = 0.049), although the production efficiency per capita female (one-way ANOVA, F3,52 = 3.576, P = 0.020) decreased at the high parasitoid density. Exposure time of more than 5 days did not appear to increase the productivity. Increased parasitoid density similarly did not appear to increase the productivity. Therefore, the combination of six pairs and 5-day exposure times seems to be most suitable for laboratory rearing.
Figure 5 shows a 6-month productivity trend of the large-scale rearing of G1 Ganaspis brasiliensis at the quarantine facility of the Edmund Mach Foundation (Trento, Italy) in 2021. The rearing was started using 150 three-day-old, mated female wasps derived from a small-scale rearing at the quarantine laboratory of CABI in Del&#233;mont, Switzerland. More wasps from the rearing were gradually added to the parasitism cage, until reaching 1,500-2,000 individuals (sex ratio 50:50) on week 5. The occupancy of the parasitism cage was thereafter maintained at the same level by adding new wasps produced in the large-scale rearing itself. During the entire period, the parasitism cage was provided with freshly infested fruit every 2-3 days. The fruit (blueberries) was offered to G1 G. brasiliensis immediately after the overnight exposure to Drosophila suzukii. Parasitoid production started 5 weeks after the initial host exposure (Figure 5A). From week 8 to week 22, the parasitoid production was proportional to the amount of fruit exposed, averaging 0.44 &#177; 0.03 g/parasitoid (mean &#177; SE; Figure 5B). A total of 53,736 parasitoids were collected, with an average female progeny of 45.9% (range: 32.4%-79.0%).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63898/63898fig01.jpg" /> 
Figure 1: A flow chart for small-scale laboratory rearing procedures of Drosophila suzukii and G3 Ganaspis brasiliensis. The left side shows the host fly rearing procedures while the right side illustrates the parasitoid rearing cycle. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63898/63898fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63898/63898fig02.jpg" /> 
Figure 2: A flow chart for the large-scale rearing procedures of Drosophila suzukii and G1 Ganaspis brasiliensis. The left side shows the host fly rearing procedures while the right side illustrates the parasitoid rearing cycle. Abbreviation: SDM = standard Drosophila medium. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63898/63898fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63898/63898fig03.jpg" /> 
Figure 3: Containers and tubes used to store and ship G1 Ganaspis brasiliensis from the large-scale rearing. (A) A horizontal view of the storage container showing a water tube inside the container, (B) a vertical view of the container showing a ventilated lid and a foam stopper, and (C) a ventilated lid and a piece of absorbent paper for (D) the shipping tube. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63898/63898fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63898/63898fig04.jpg" /> 
Figure 4: Representative example for small-scale laboratory rearing of G3 Ganaspis brasiliensis. (A) Number of offspring produced per flask, (B) number of offspring produced per female wasp, and (C) percentage of female offspring, under two different parasitoid densities and exposure times. Values are mean &#177; SE, and bars bearing different letters are significantly different (ANOVA, Tukey&#39;s HSD, P &#60; 0.05). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63898/63898fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63898/63898fig05.jpg" /> 
Figure 5: Productivity trend of the large-scale rearing from its onset through week 23. (A) Bars indicate the numbers of G1 Ganaspis brasiliensis offspring collected weekly from the eclosion cages to replace old individuals in the parasitism cage (dark green) and to be stored or shipped (light green). The orange line indicates the amount of host-infested fruit (kg of blueberries) provided each week to the parasitoids within the parasitism cage. (B) Weekly ratio of the weight of host-infested fruit to the number of parasitoid offspring produced 5 weeks after exposure (i.e., grams of fruit required to produce a single adult parasitoid). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63898/63898fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Methods for Rearing the Parasitoid Ganaspis brasiliensis, a Promising Biological Control Agent for the Invasive Drosophila suzukii at JoVE.com

Methods for Rearing the Parasitoid Ganaspis brasiliensis, a Promising Biological Control Agent for the Invasive Drosophila suzukii

Ganaspis brasiliensis-a larval parasitoid of Drosophila suzukii (a global invasive fruit crop pest)-has been approved or is considered for introduction into Europe and the United States for biological control of this pest. This article provides protocols for both small-scale and large-scale rearing of this parasitoid.

Methods for Rearing the Parasitoid Ganaspis brasiliensis, a Promising Biological Control Agent for the Invasive Drosophila suzukii

Native to East Asia, the spotted-wing drosophila, Drosophila suzukii (Matsumura) (Diptera: Drosophilidae), has established widely in the Americas, Europe, ...

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4.1K Views.  Fondazione Edmund Mach. Langdurig onderzoek en gebruik van een biocontrolemiddel zijn afhankelijk van de beschikbaarheid van effectieve en economische opfoktechnieken, en dit protocol heeft bewezen efficiënt te zijn voor het fokken van deze parasitoïde. Er is momenteel enorme interesse in het fokken van deze parasitoïde voor onderzoek of veldontlading, en het protocol zal toekomstig langetermijnonderzoek en gebruik van dit veelbelovende biocontrolemiddel ten goede komen. Eerst moeten gebruikers mogelijk een basis...

Video: Methods for Rearing the Parasitoid Ganaspis brasiliensis, a Promising Biological Control Agent for the Invasive Drosophila suzukii

Mass Production of Genetically Modified Aedes aegypti for Field Releases in Brazil

New techniques and methods are being sought to try to win the battle against mosquitoes. Recent advances in molecular techniques have led to the development of new and innovative methods of mosquito control based around the Sterile Insect Technique (SIT)1-3. A control method known as RIDL&#160;(Release of Insects carrying a Dominant Lethal)4, is based around SIT, but uses genetic methods to remove the need for radiation-sterilization5-8. A RIDL&#160;strain of Ae. aegypti was successfully tested in the field in Grand Cayman9,10; further field use is planned or in progress in other countries around the world.
Mass rearing of insects has been established in several insect species and to levels of billions a week. However, in mosquitoes, rearing has generally been performed on a much smaller scale, with most large scale rearing being performed in the 1970s and 80s. For a RIDL&#160;program it is desirable to release as few females as possible as they bite and transmit disease. In a mass rearing program&#160;there are several stages to produce the males to be released: egg production, rearing eggs until pupation, and then sorting males from females before release. These males are then used for a RIDL&#160;control program,&#160;released as either pupae or adults11,12.
To suppress a mosquito population using RIDL&#160;a large number of high quality male adults need to be reared13,14. The following describes the methods for the mass rearing of OX513A, a RIDL&#160;strain of Ae. aegypti 8, for release and covers the techniques required for the production of eggs and mass rearing RIDL&#160;males for a control program.

Mass Production of Genetically Modified Aedes aegypti for Field Releases in Brazil

To achieve population suppression of Aedes aegypti using the RIDL&#174; (Release of Insects carrying a Dominant Lethal) system, large numbers of male mosquitoes need to be released. This requires the use of mass rearing techniques and technology to provide reliable systems to obtain the maximum number of high quality male mosquitoes.

New techniques and methods are being sought to try to win the battle against mosquitoes. Recent advances in molecular techniques have led to the ...

Environment

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is a model for understanding microbial interactions with animal hosts, facilitated by approaches to rear large sample sizes of Drosophila under microorganism-free (axenic) conditions, or with defined microbial communities (gnotobiotic). This work outlines a method for collection of Drosophila embryos, hypochlorite dechorionation and sterilization, and transfer to sterile diet. Sterilized embryos are transferred to sterile diet in 50 ml centrifuge tubes, and developing larvae and adults remain free of any exogenous microbes until the vials are opened. Alternatively, flies with a defined microbiota can be reared by inoculating sterile diet and embryos with microbial species of interest. We describe the introduction of 4 bacterial species to establish a representative gnotobiotic microbiota in Drosophila. Finally, we describe approaches for confirming bacterial community composition, including testing if axenic Drosophila remain bacteria-free into adulthood.

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

A method for rearing Drosophila melanogaster under axenic and gnotobiotic conditions is presented. Fly embryos are dechorionated in sodium hypochlorite, transferred aseptically to sterile diet, and reared in closed containers. Inoculating diet and embryos with bacteria leads to gnotobiotic associations, and bacterial presence is confirmed by plating whole-body Drosophila homogenates.

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is ...

Developmental Biology

Flying Insect Detection and Classification with Inexpensive Sensors

An inexpensive, noninvasive system that could accurately classify flying insects would have important implications for entomological research, and allow for the development of many useful applications in vector and pest control for both medical and agricultural entomology. Given this, the last sixty years have seen many research efforts devoted to this task. To date, however, none of this research has had a lasting impact. In this work, we show that pseudo-acoustic optical sensors can produce superior data; that additional features, both intrinsic and extrinsic to the insect&#8217;s flight behavior, can be exploited to improve insect classification; that a Bayesian classification approach allows to efficiently learn classification models that are very robust to over-fitting, and a general classification framework allows to easily incorporate arbitrary number of features. We demonstrate the findings with large-scale experiments that dwarf all previous works combined, as measured by the number of insects and the number of species considered.

We proposed a system that uses inexpensive, noninvasive pseudo-acoustic optical sensors to automatically and accurately detect, count, and classify flying insects based on their flying sound.

An inexpensive, noninvasive system that could accurately classify flying insects would have important implications for entomological research, and allow ...

Bioengineering

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, Drosophila research lacks popularity in developing countries, possibly due to the misinformed idea that establishing a lab and performing relevant experiments with such tiny insects is difficult and requires expensive, specialized apparatuses. Here, we describe how to build an affordable flylab to quantitatively analyze a myriad of behavioral parameters in D. melanogaster, by 3D-printing many of the necessary pieces of equipment. We provide protocols to build in-house vial racks, courtship arenas, apparatuses for locomotor assays, etc., to be used for general fly maintenance and to perform behavioral experiments using adult flies and larvae. We also provide protocols on how to use more sophisticated systems, such as a high resolution oxygraph, to measure mitochondrial oxygen consumption in larval samples, and show its association with behavioral changes in the larvae upon the xenotopic expression of the mitochondrial alternative oxidase (AOX). AOX increases larval activity and mitochondrial leak respiration, and accelerates development at low temperatures, which is consistent with a thermogenic role for the enzyme. We hope these protocols will inspire researchers, especially from developing countries, to use Drosophila to easily combine behavior and mitochondrial metabolism data, which may lead to information on genes and/or environmental conditions that may also regulate human physiology and disease states.

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

We provide protocols for anyone with a "maker culture" mind to start building a flylab for quantitative analysis of a myriad of behavioral parameters in Drosophila melanogaster, by 3D-printing many of the necessary pieces of equipment. We also describe a high resolution respirometry protocol using larvae to combine behavioral and mitochondrial metabolism data.

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, ...

Behavior

Electroporation-mediated RNA Interference Method in Odonata

Dragonflies and damselflies (order Odonata) represent one of the most ancestral insects with metamorphosis, in which they change their habitat, morphology, and behavior drastically from aquatic larvae to terrestrial/aerial adults without pupal stage. Odonata adults have a well-developed color vision and show a remarkable diversity in body colors and patterns across sexes, stages, and species. While many ecological and behavioral studies on Odonata have been conducted, molecular genetic studies have been scarce mainly due to the difficulty in applying gene functional analysis to Odonata. For instance, RNA interference (RNAi) is less effective in the Odonata, as reported in the Lepidoptera. To overcome this problem, we successfully established an RNAi method combined with in vivo electroporation. Here we provide a detailed protocol including a video of the electroporation-mediated RNAi method as follows: preparation of larvae, species identification, preparation of dsRNA/siRNA solution and injection needles, ice-cold anesthesia of larvae, dsRNA/siRNA injection, in vivo electroporation, and individual rearing until adult emergence. The electroporation-mediated RNAi method is applicable to both damselflies (suborder Zygoptera) and dragonflies (suborder Anisoptera). In this protocol, we present the methods for the blue-tailed damselfly Ischnura senegalensis (Coenagrionidae) as an example of damselfly species and the pied skimmer dragonfly Pseudothemis zonata (Libellulidae) as another example of dragonfly species. As representative examples, we show the results of RNAi targeting the melanin synthesis gene multicopper oxidase 2. This RNAi method will facilitate understanding of various gene functions involved in metamorphosis, morphogenesis, color pattern formation, and other biological features of Odonata. Moreover, this protocol may be generally applicable to non-model organisms in which RNAi is less effective in gene suppression due to the inefficiency and low penetrance.

We provide a detailed protocol for electroporation-mediated RNA interference in insects of the order Odonata (dragonflies and damselflies) using the blue-tailed damselfly (Ischnura senegalensis: Coenagironidae: Zygoptera) and the pied skimmer dragonfly (Pseudothemis zonata: Libellulidae: Anisoptera).

Dragonflies and damselflies (order Odonata) represent one of the most ancestral insects with metamorphosis, in which they change their habitat, ...

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Fopius arisanus (Sonan) is an important parasitoid of Tephritid fruit flies for at least two reasons. First, it is the one of only three opiine parasitoids known to infect the host during the egg stage1. Second, it has a wide range of potential fruit fly hosts. Perhaps due to its life history, F. arisanus has been a successfully used for biological control of fruit flies in multiple tropical regions2-4. One impediment to the wide use of F. arisanus for fruit fly control is that it is difficult to establish a stable laboratory colony5-9. Despite this difficulty, in the 1990s USDA researchers developed a reliable method to maintain laboratory populations of F. arisanus10-12. There is significant interest in F. arisanus biology13,14, especially regarding its ability to colonize a wide variety of Tephritid hosts14-17; interest is especially driven by the alarming spread of Bactrocera fruit fly pests to new continents in the last decade18. Further research on F. arisanus and additional deployments of this species as a biological control agent will benefit from optimizations and improvements of rearing methods. In this protocol and associated video article we describe an optimized method for rearing F. arisanus based on a previously described approach12. The method we describe here allows rearing of F. arisanus in a small scale without the use of fruit, using materials available in tropical regions around the world and with relatively low manual labor requirements.

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Fopius arisanus is an egg-larval parasitoid of Tephritid fruit flies that is successfully used in biological control of these important tropical pests. We describe here an optimized protocol for rearing F. arisanus on a small scale using readily available materials.

Fopius arisanus (Sonan) is an important parasitoid of Tephritid fruit flies for at least two reasons. First, it is the one of only three opiine ...

Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory

Emergent properties and external factors (population-level and ecosystem-level interactions, in particular) play important roles in mediating ecologically-important endpoints, though they are rarely considered in toxicological studies. D. melanogaster is emerging as a toxicology model for the behavioral, neurological, and genetic impacts of toxicants, to name a few. More importantly, species in the genus Drosophila can be utilized as a model system for an integrative framework approach to incorporate emergent properties and answer ecologically-relevant questions in&#160;toxicology research. The aim of this paper is to provide a protocol for exposing species in the genus Drosophila to pollutants to be used as a model system for a range of phenotypic outputs and ecologically-relevant questions. More specifically, this protocol can be used to 1) link multiple biological levels of organization and understand the impact of toxicants on both individual- and population-level fitness; 2) test the impact of toxicants at different stages of developmental exposure; 3) test multigenerational and evolutionary implications of pollutants; and 4) test multiple contaminants and stressors simultaneously.

Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory

In this paper, we provide a detailed protocol for exposing species in the genus Drosophila to pollutants with the goal of studying the impact of exposure on a range of phenotypic outputs at different developmental stages and for more than one generation.

Emergent properties and external factors (population-level and ecosystem-level interactions, in particular) play important roles in mediating ...

Simple Homemade Tools to Handle Fruit Flies&#8212;Drosophila melanogaster

The fruit fly, Drosophila melanogaster, is widely used both in biological research and biology education. Handling adult flies is common but difficult in practice, as adult flies fly. Demonstrated here is how to make some simple and cost-effective tools to address difficult issues in the handling of Drosophila. Holes in foam stoppers are made and pipette tips or funnels are inserted into the holes. Flies then move only in one direction into the pipette tip/funnel assemblage, allowing efficient control of the transfer of adult Drosophila into or out of a vial. Existing protocols have been modified for cool-anesthetizing flies by chilling in crushed ice and transferring them onto a cold, hard icepack surface. The icepack is covered with a piece of medical gauze that keeps immobilized flies from the condensed water when examined under a stereomicroscope. The flies are finally euthanized for counting and sorting or discarded by microwaving. A bottle-shaped cage has also been developed for collecting eggs, as well as a labor-saving device and accompanying protocol for cleaning glass culture vials.

Described here is the use of several homemade tools to transfer, chill, and kill adult Drosophila, as well as to clean glass culture vials and collect eggs. These tools are easy to make and are rather efficient in handling Drosophila.

The fruit fly, Drosophila melanogaster, is widely used both in biological research and biology education. Handling adult flies is common but difficult in ...

Topical Application Bioassay to Quantify Insecticide Toxicity for Mosquitoes and Fruit Flies

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. Insecticide resistance surveillance of mosquito populations is typically done through Centers for Disease Control and Prevention (CDC) bottle bioassays or World Health Organization (WHO) tube tests. However, these methods can result in a high degree of variability in mortality data due to variable insecticide contact with&#160;the insect, the relatively small numbers of organisms tested, extensive variation in mass between populations, and constantly changing environmental conditions, leading to variable outcomes. This paper presents the topical application bioassay, adapted as a high-throughput phenotypic bioassay for both mosquitoes and fruit flies, to test large numbers of insects along&#160;a range of insecticide concentrations.
This assay 1) ensures consistent treatment and insecticide contact with every organism, 2) produces highly specific dose-response curves that account for differences in average mass between strains and sexes (which is particularly important for field-collected organisms), and 3) allows for the calculation of statistically rigorous median lethal doses (LD50), which are necessary for resistance ratio comparisons-an alternative surveillance approach from diagnostic dose mortality, which is also used for larvicide resistance surveillance. This assay will be a complementary tool for accurately phenotyping mosquito populations and, as illustrated using fruit flies, is easily adaptable for use with other insects. We argue that this assay will help fill the gap between genotypic and phenotypic insecticide resistance in multiple insect species.

We describe the methodology and importance of the topical application bioassay to measure insecticide susceptibility in mosquitoes and fruit flies. The presented assay is high-throughput, utilizes insect mass-thus allowing for calculating a mass-relativized lethal dose instead of concentration-and likely has lower variability than other similar methods.

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. ...

Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis

The Oriental fruit fly, Bactrocera dorsalis, is a highly invasive and adaptive pest species that causes damage to citrus and over 150 other fruit crops worldwide. Since adult fruit flies have great flight capacity and females lay their eggs under the skins of fruit, insecticides requiring direct contact with the pest usually perform poorly in the field. With the development of molecular biological tools and high-throughput sequencing technology, many scientists are attempting to develop environmentally friendly pest management strategies. These include RNAi or gene editing-based pesticides that downregulate or silence genes (molecular targets), such as olfactory genes involved in searching behavior, in various insect pests. To adapt these strategies for Oriental fruit fly control, effective methods for functional gene research are needed. Genes with critical functions in the survival and reproduction of B. dorsalis serve as good molecular targets for gene knockdown and/or silencing. The CRISPR/Cas9 system is a reliable technique used for gene editing, especially in insects. This paper presents a systematic method for CRISPR/Cas9 mutagenesis of B. dorsalis, including the design and synthesis of guide RNAs, collecting embryos, embryo injection, insect rearing, and mutant screening. These protocols will serve as a useful guide for generating mutant flies for researchers interested in functional gene studies in B. dorsalis.

Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis

This paper presents the step-by-step protocols for CRISPR/Cas9 mutagenesis of the Oriental fruit fly Bactrocera dorsalis. Detailed steps provided by this standardized protocol will serve as a useful guide for generating mutant flies for functional gene studies in B. dorsalis.

The Oriental fruit fly, Bactrocera dorsalis, is a highly invasive and adaptive pest species that causes damage to citrus and over 150 other fruit crops ...

Methods for Rearing the Parasitoid Ganaspis brasiliensis, a Promising Biological Control Agent for the Invasive Drosophila suzukii

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