Protocols for Efficient Rearing and Functional Analysis of the Endoparasitoid Wasp Asobara japonica and its Host Drosophila melanogaster

Summary

The goal of this protocol is to facilitate functional analysis of parasitoid-host interactions in Asobara japonica and Drosophila melanogaster through optimized rearing methods, a single-oviposition infection assay, and RNAi-mediated gene knockdown.

Abstract

Parasitism is a biological interaction in which one organism exploits the body or resources of another (the host), resulting in substantial damage or death to the host. Among parasitic animals, parasitoid wasps are one of the most species-rich lineages, accounting for nearly 20% of all insect species. In particular, endoparasitoid wasps oviposit directly into host bodies and deploy a diverse array of venom factors to manipulate host development, immunity, and physiology. In response to wasp attack, hosts attempt to eliminate parasitoid eggs through innate immune mechanisms. However, the molecular and cellular mechanisms by which individual venom components modulate host biology and promote successful parasitism remain poorly understood. Here, we describe a standardized laboratory protocol for rearing the endoparasitoid wasp Asobara japonica and its host Drosophila melanogaster. Both parthenogenetic and sexual strains of A. japonica are available, and the parthenogenetic strain exhibits a high parasitism success rate, allowing stable maintenance of laboratory stocks for genome analysis and parasitism assays. A single-oviposition infection assay and a double-stranded RNA-based gene knockdown method were optimized for functional analysis of venom genes. Together, these protocols provide a practical experimental framework for dissecting the molecular mechanisms underlying parasitoid-host interactions and will facilitate future research in developmental biology, immunology, and physiology.

Introduction

Parasitism is a lifestyle in which one organism exploits the nutritional resources of another and is widespread across taxa. It is estimated that up to half of all known organisms exhibit parasitism in their life cycle¹. Among parasites, parasitoid wasps are a particularly diverse group, accounting for approximately 20% of all insects². They exploit a wide range of arthropod hosts, such as insects, spiders, and mites, and exhibit remarkable diversity in their life histories and parasitic strategies³. These features make parasitoid wasps one of the most evolutionarily and ecologically successful insect groups.

Interactions between parasitoids and their hosts have been extensively studied as model systems of evolutionary “arms races.” In particular, endoparasitoid wasps deploy a diverse array of factors, including venoms, symbiotic viruses, and teratocytes^4,5. These factors manipulate host physiology and development, thereby ensuring successful development within the host. Despite these advances, the molecular mechanisms underlying parasitism remain poorly understood across most parasitoid species. One major limitation is their small body size, which has hindered the biochemical identification of venom components. In addition, it is technically challenging to maintain parasitoids and their hosts with synchronized developmental stages under laboratory conditions.

Recent advances in next-generation sequencing and other omics technologies now enable genome-wide analyses even in very small parasitoid species. These approaches have opened new avenues for investigating previously inaccessible molecular mechanisms, allowing parasitoid-host interactions to be studied at unprecedented resolution. In this context, our laboratory focuses on Asobara japonica Belokobylskij (Hymenoptera: Braconidae), a species originally identified in Japan^6,7,8. A. japonica includes a thelytokous parthenogenetic strain, in which females produce offspring without mating. This feature allows us to collect substantial amounts of genetically uniform genomic DNA for whole-genome sequencing⁹. In addition, this species parasitizes a broad range of Drosophila species, including the model organism D. melanogaster, which offers major experimental advantages due to its well-established rearing conditions and precisely defined developmental stages, enabling reproducible infection assays and accurate control of parasitoid development¹⁰.

During parasitism of A. japonica, a female wasp injects venom together with a single egg into a fly larva, inside which a wasp larva grows alongside its host. Notably, A. japonica exhibits a high parasitism success rate on the host D. melanogaster, making this parasitoid-host pair a powerful experimental model. Here, we describe protocols for stable rearing, a single-oviposition infection assay, and a double-stranded RNA (dsRNA)-based gene knockdown method in A. japonica. These protocols provide a reliable framework for dissecting molecular mechanisms underlying parasitoid-host interactions.

Protocol

A schematic overview of the synchronized rearing method for D. melanogaster and A. japonica is presented in Figure 1.

1. Rearing the host fruit fly Drosophila melanogaster in the laboratory

1. Maintain all experimental animals at 25 °C and 50-60% relative humidity under a 12:12 h light/dark cycle. Use the wild-type strain Oregon-R (OR) as a host for A. japonica (Figure 2A).
2. Transfer adult flies to a new food vial containing standard cornmeal-yeast-agar medium (0.55% agar, Table 1). Label the vial with the date of transfer and the strain name "OR".
3. Clear these adult flies one day after the transfer to prevent an excessive number of eggs in the food. Make sure that offspring larvae are feeding on the surface of the food.
   NOTE: As a guideline, maintain approximately 200-250 eggs per food vial (φ30×100 mm). Higher egg densities may reduce parasitism success and increase variation in host and wasp development.
4. Four days after Step 1.2, transfer a spoonful of fly larvae (4 days after egg laying, dAEL) to a new food vial. Keep these larvae unexposed to parasitoids for the next generation. This procedure also helps to control the host larval density in a vial.

2. Rearing the endoparasitoid wasp Asobara japonica in the laboratory

1. Use a parthenogenetic strain Tokyo (TK) and a sexual strain Iriomote-jima (IR) (Figure 2B). The former consists of females, whereas the latter consists of females and males. Males eclose two-four days earlier than females. Therefore, confirm the presence of females by their ovipositors before the transfer.
2. Transfer 10-20 adult wasps directly from the stock vial into the vial containing fly larvae at 4 dAEL. Label the vial with the date of transfer and the strain name "TK" or "IR". Maintain the wasps in the same conditions as Step 1.1.
3. Allow adult wasps to oviposit in fly larvae. Clear these wasps one to two days later to prevent superparasitism, which can lead to host death. Remove eclosed adult flies that escaped from parasitism 10 or more days after Step 2.2.
4. Two weeks after Step 2.2, adult wasps begin to eclose from the host fly pupal case. Transfer these newly-eclosed adult wasps to a new food vial with filter paper containing 25% glucose and 50 mM Vitamin C solution. The pieces of filter paper should be sterilized at 130 °C for 2 h. Providing the solution on filter paper promotes oviposition behaviors and prolongs the lifespan of adult wasps. Our approach to using vitamin C to enhance oviposition was inspired by observations in Drosophila studies¹¹.

3. Single-Oviposition Infection Assay

1. Synchronize the developmental stage of fly larvae used for the infection experiment.
   1. Place adult flies in a 50-mL tube containing a grape-juice agar plate (3% agar) spread with fresh yeast paste and allow them to lay eggs for 24 h.
   2. Collect newly-hatched first-instar larvae on the grape-juice agar plate and transfer them to a small vial (12 mL) containing 2-3 g fly food paste (fly food mixed with an appropriate amount of water). 20-30 larvae are placed in the vial. The fly larvae are allowed to grow at 25 °C.
2. At the appropriate time point, prepare a 4 cm Petri dish containing a thin, evenly spread layer of fly food paste covering an area approximately 1.5-2.0 cm in diameter (hereafter referred to as infection assay arena). In addition to the infection assay arena, prepare a new small vial (12 mL) containing an appropriate amount of fly food paste. This vial is used for rearing the host larvae after the infection assay.
3. Collect staged larvae and transfer them to a dish containing deionized water (resistivity = 15 MΩ·cm, 25 °C). Rinse the larvae in the dish with gentle agitation to remove residual food from their bodies.
4. After rinsing, gently remove excess moisture from the larvae using lint-free paper wipes, as excess water interferes with wasp oviposition behavior on the infection assay arena. Then transfer the larvae to the arena. This step increases larval activity during the infection experiment.
5. Anesthetize adult wasps with CO₂ gas at a flow rate of 5 L·min^-1 and a pressure of 0.2 MPa. Then, gently place approximately three female wasps into the infection assay arena. Close the lid of the dish and allow them to recover on fly food paste.
6. Observe the oviposition behavior of these wasps under a stereomicroscope (zoom setting: 10× to 15×).
   NOTE: When a wasp touches a moving fly larval body, it quickly inserts its ovipositor into the fly body and injects venom components. Although the fly larva writhes and thrashes its body, it immediately becomes paralyzed. Subsequently, the wasp vibrates its abdomen and deposits an egg into the host larval body, after which the ovipositor is withdrawn. This oviposition behavior is referred to as infection.
7. After infection, open the lid and transfer the infected fly larva to the new small vial containing fly food paste prepared in Step 3.2. To standardize infection timing, perform the infection experiment within 15 min.
8. Label the small vial with experimental information, such as the date, sample name, infection time, and number of individuals. These vials are plugged and then placed in a humid container and kept at 25 ˚C.

4. Generation of RNA interference (RNAi) wasps by dsRNA injection

1. Synthesize dsRNAs of 400 bp or longer for dsRNA-mediated gene knockdown (RNA interference, RNAi) using an in vitro transcription-based dsRNA synthesis system. The dsRNA samples should be kept at -20 to -80 °C until use. Detailed protocols are as previously described⁹.
2. Prepare a 5.5 cm petri dish containing 2% agar for placing wasp bodies. Fill two-thirds of each dish with agar to generate a thick layer of agar plate for injection.
3. Place a lint-free paper wipe moistened with water on the lid of a plastic dish.
4. Collect host fly pupae at 7 days post-infection (dpi) from the vial and place them on the lint-free paper wipes prepared in Step 4.3. Adding a drop of water to the vial wall helps to detach host fly pupal cases from the vial.
5. Under the stereomicroscope (zoom setting: 20×), remove the host fly pupal case and expose a wasp body with forceps. At 7 dpi, most wasps are in the pupal stage, in which the head, thorax, and abdomen parts are distinguishable. A wasp body is immobile.
6. Place the retrieved wasp bodies on the agar plate prepared in Step 4.2, aligning their body axes in the same orientation. Arrange 20-30 wasp bodies with their dorsal side up in one to two lines. Remove excess moisture on the agar plate using a lint-free paper wipe to prevent wasps from slipping.
7. Following the manufacturer’s instructions, set up the injection apparatus and stereomicroscope (Figure 3A). Use a microinjection system equipped with a glass capillary needle that allows precise control of injection volume and injection speed. Place the agar plate in which the wasp heads are oriented toward the tip of the injection needle. The injection volume is 50 nL·wasp^-1 and the injection speed is 50 nL·sec^-1 on the injector.
8. Prepare glass capillary needles using a needle puller. The shape of glass capillary is adjustable with heater temperature.
9. Place the glass capillary needle on the needle grinder at an angle of 30-35°. Grind the needle until the tip length reaches 0.04 mm (four divisions on the eyepiece reticle, Figure 3B).
10. Just before injection, thaw dsRNA. The concentration of dsRNA is 0.25-1 µg·µL^-1. For visualization, the dsRNA solution is colored with a blue dye (Erioglaucine disodium salt, 2% weight·volume^-1). Using a microloader pipette tip, fill the glass capillary with dsRNA solution with blue dye. 4 µL is sufficient for injecting 30 wasp bodies. Ensure that no air bubbles are present inside the solution.
    NOTE: After loading the dsRNA solution, gently tap the side of the needle with a fingernail to remove any trapped air bubbles. Residual air bubbles can obstruct the flow of the solution and interfere with successful injection.
11. Using a 1 mL syringe and a needle, fill the unoccupied portion of the glass capillary with mineral oil, ensuring that no air bubbles remain inside the capillary.
12. Attach the glass capillary needle containing dsRNA solution and mineral oil to the needle holder and then mount it onto the injection device.
13. Adjust the position of the agar plate and the glass capillary needle on the injector, so that the glass needle is positioned at a 30-45° angle relative to the wasp bodies (Figure 3A).
14. Insert the glass capillary needle into the dorsal side of the wasp body, with the slightly lateral side to the dorsal midline. Microinject the dsRNA solution inside the body. The injected dsRNA solution is visible with blue dye (Figure 3C).
15. After the injection, place the agar plates in a humid container (70-80% relative humidity) and maintain them in a 25 °C incubator for a week, until adult wasps eclose. Injured wasp bodies should be removed from the agar plate to prevent bacterial contamination.
    PAUSE POINT: In Step 4.6, the wasp pupae can be kept on the agar plate for several hours before injection. Needles can be prepared in advance (Step 4.8 and 4.9); however, they are prone to clogging over time. Freshly prepared needles generally provide more reliable injection performance.

Results

The asexual and sexual strains of Asobara japonica
A. japonica is widely distributed throughout Japan. Thelytokous parthenogenetic strains are predominant in the main islands, while arrhenotokous sexual strains are found in the subtropical islands, including Amami-Oshima and Iriomote-jima islands^8,12. In sexual strains, males emerge earlier than females by a few days so that the timing of adult transfer is distinct between these strains. Thelytokous parthenogenesis in A. japonica is induced by infection with Wolbachia, a maternally transmitted endosymbiotic bacterium¹³. Elimination of Wolbachia by antibiotic treatment results in the production of males^9,14, and transplantation of Wolbachia into sexual strains induces parthenogenesis¹⁵. Both asexual and sexual strains of A. japonica exhibit high parasitism success across multiple Drosophila species. For example, they can parasitize D. suzukii, an economically important fruit pest, and are considered potential biological control agents in agriculture¹⁶. Among the available strains, we primarily used the Wolbachia-infected TK strain for our experiments. The TK strain exhibits higher parasitism activity toward host larvae compared to the IR strain¹⁴, making it suitable for analyses.

Parasitism success depends on rearing conditions
Parasitism success rate of A. japonica is generally high, but it is strongly affected by the rearing condition of host flies. Under nutrient-poor conditions, the growth period of fly larvae is prolonged and the timing of pupariation is delayed¹⁷. Under these conditions, wasps may fail to properly utilize host resources and to synchronize their development with that of hosts. To examine this possibility, we reared flies and wasps under normal-yeast (1×) and yeast-poor (0.1×) conditions. We then evaluated parasitism success and developmental timing (Figures 2C, 2D). Parasitism success was calculated as the ratio of emerged adult wasps to host larvae.

Under normal-yeast conditions, the parasitism success rate was 81.7% in the single-oviposition infection assay (Figure 2C, left; Table 2). When the yeast content was reduced to 10% (yeast-poor condition), the parasitism success rate decreased to 35.0% (Figure 2C, right; Table 2). This reduction may be explained by the fact that approximately 50% of host larvae died following infection. These results suggest that resource allocation between host and parasitoid is disrupted under nutrient-poor conditions.

Notably, the timing of wasp eclosion was not delayed to the same extent under yeast-poor conditions as it was under normal-yeast conditions. While fly eclosion timing was delayed by up to 9 days, wasp eclosion timing was delayed by only up to 4-5 days. This result suggests that nutrient availability primarily affects host development. The timing of eclosion is consistent both in flies and wasps under normal-yeast conditions, but not yeast-poor conditions, suggesting that synchronized development between parasitoids and hosts depends on the nutrient availability.

RNAi-based approach for functional analysis of venom genes
Although transcriptomic and proteomic analyses of the venom gland identify numerous candidate molecules, it remains unclear which of these are required for successful parasitism. To determine the physiological roles of these candidate molecules, in vivo functional analyses using gene knockdown approaches are required.

We recently reported that A. japonica parasitism induces apoptosis, autophagy, and mitotic arrest in the imaginal discs of D. melanogaster larvae, resulting in severe tissue degradation, a phenomenon termed imaginal disc degradation (IDD)¹⁰. To identify the molecules responsible for IDD, we performed RNAi screen of candidate genes identified from transcriptomic, proteomic, and comparative genomic analyses. As a result, two genes, imaginal disc degradation factor (IDDF)-1 and IDDF-2 were identified. Both IDDF-1 and IDDF-2 contain the Domain of Unknown Function 4803 (DUF4803). Notably, following our initial report, an independent study identified multiple A. japonica venom proteins (termed Venom-Induced Degradation, VID) associated with IDD, all of which also possess the DUF4803 domain¹⁸.

Based on these results, we further optimized the RNAi protocol to improve reproducibility and efficiency. In particular, the use of Nanoject III enabled precise control of injection volume and speed, resulting in more consistent knockdown outcomes.

To evaluate the efficiency of the optimized RNAi protocol, we performed knockdown of IDDF-1. In eclosed adult wasps, the expression level of IDDF-1 was significantly decreased compared with the control (Figure 3D). For functional validation, we performed single-oviposition infection assays using these RNAi wasps and host fly larvae expressing the apoptosis reporter GC3Ai in the wing pouch region¹⁹. A GC3Ai signal was detected in the wing pouch following infection with control (GFP RNAi) wasps. In contrast, the GC3Ai signal was significantly reduced following infection with IDDF-1 RNAi wasps (Figure 3E, 3F). These results are consistent with previous findings, confirming that IDDF-1 function is effectively suppressed by RNAi.

Figure 1. Schematic overview of the rearing procedures for D. melanogaster and A. japonica. 
Step-wise procedures are shown according to the developmental stages of D. melanogaster. Please click here to view a larger version of this figure.

Figure 2. Parasitism by Asobara japonica. (A) Schematic of the developmental relationship between D. melanogaster and its parasitoid A. japonica. A female wasp deposits venoms and a single egg into a fly larva. After hatching, the wasp larva develops in parallel with the host larva, which subsequently pupariates. Following pupariation, the wasp consumes the whole fly body and eventually ecloses from the fly pupal case. (B) Adult wasps of the asexual strain (left) and sexual strain (right) of A. japonica. Scale bar, 1 mm. (C) Emergence rate of flies or wasps under normal (1×) and yeast-poor (0.1×) conditions. Single-oviposition infection assays were performed using host fly larvae at 4 days after egg laying (dAEL). n = 267 (1×, flies), 120 (0.1×, flies), 85 (1×, wasps), and 80 (0.1×, wasps), respectively. (D) Eclosion timing of flies and wasps under the two nutritional conditions shown in (C). The ratio of adults to pupae are indicated. n = 250, 107, 61, and 41, respectively. The black arrow indicates the timing of wasp infection. Please click here to view a larger version of this figure.

Figure 3. Optimized RNAi procedure using Nanoject III. (A) Setup of the microinjection apparatus under a stereomicroscope. The glass capillary needle was positioned at an angle of 30-45° relative to the agar plate. (B) The tip of the glass capillary needle was sharpened using a needle grinder. (C) A blue-dyed solution was microinjected into a wasp body using a glass capillary needle. The anterior side of the wasp is indicated by A in the image. P, posterior. Scale bar, 1 mm. (D) Expression levels of IDDF-1 in GFP RNAi and IDDF-1 RNAi wasps were quantified using the delta-delta C_t method. Expression levels were normalized to RpL32-Aj. Data represent the mean ± SD with individual data points (n = 5). ***P < 0.005 (Student’s t-test). (E and F) GC3Ai signals in wing discs of uninfected larvae and host larvae at 6 h post-infection (hpi) following infection with control (GFP RNAi) or IDDF-1 RNAi wasps. Wing discs are outlined with white dotted lines based on DAPI staining of nuclei (1:10000, 62247; Thermo Fisher Scientific). The ratio of GC3Ai-positive area to wing pouch area was quantified. ***P < 0.005 (One-way ANOVA followed by Tukey’s multiple comparisons test). n = 11 biological replicates per group. Scale bar, 100 µm. Please click here to view a larger version of this figure.

Components	Amount
Water	1 L
Agar	5.5 g
Yeast	40 g
Corn meal	90 g
Glucose	100 g
Propionic acid	3 mL
10% butyl p-Hydroxybenzoate in 70% EtOH	3.5 mL

Table 1: Components of standard cornmeal-yeast-agar medium (Fly food).

Food type	Condition	# of total individuals	# of eclosed or dead individuals				Emergence rate of flies or wasps (%)
			Fly	Wasp	Pupal lethal	Larval lethal
1×	Uninfected	267	248	0	2	17	92.88389513
	A. japonica infected	120	0	98	9	13	81.66666667
0.1×	Uninfected	85	38	0	23	24	44.70588235
	A. japonica infected	80	0	28	13	39	35

Table 2: Number of individuals and emergence rates under normal (1x) and yeast-poor (0.1x) conditions.

Primer name	Sequence (5'>3')	Purpose
AjRpL32_qPCR_Fwd	CCCGTCACATGCTTCCTACT	qRT-PCR
AjRpL32_qPCR_Rev	GAATTTGCGATTCTGCATCA	qRT-PCR
qPCR_gene007424_Fwd	TGGGCAATGATTGTGTGAGT	qRT-PCR
qPCR_gene007424_Rev	GCGTCCCTGTATCGAAGGTA	qRT-PCR
Aj_gene007424_ex1_Fwd	ATGAGCATCAAGCAAGCAGTGCTAG	CDS cloning
Aj_gene007424_ex10_Rev	TCAGCACCTTCTACCCGGCAAATT	CDS cloning, dsRNA template
Aj_gene007424_RNAi_Fwd	GAGGAGAGTGGAAGCCAGTTGTAG	dsRNA template
T7_Aj_gene007424_ex10_Rev	GGATCCTAATACGACTCACTATAGGT CAGCACCTTCTACCCGGCAAATT	dsRNA template
T7_Aj_gene007424_ RNAi_Fwd	GGATCCTAATACGACTCACTATAGGGAG GAGAGTGGAAGCCAGTTGTAG	dsRNA template
※ These primers and dsRNA templates are reported in Kamiyama et al. 2022.
※ gene007424 stands for IDDF-1.

Table 3: Sequences of primers and dsRNA templates used in this study.

Discussion

In this study, we established a versatile experimental platform for investigating parasitoid-host interactions at the molecular level. By optimizing rearing conditions, a single-oviposition infection assay, and the RNAi protocol, we provide a robust system for functional studies of parasitoid-derived factors. This platform will facilitate future studies aimed at uncovering the mechanisms by which parasitoid wasps manipulate host development and may also contribute to the development of biological pest control strategies.

The Wolbachia-infected TK strain of A. japonica provides a highly reproducible experimental system for analyzing parasitism. In many parasitoid-host studies, experimental outcomes are often variable, with host survival and parasitoid success depending sensitively on environmental and physiological conditions. In contrast, the TK strain exhibits consistently high parasitism success, effectively creating a “parasitoid-dominant” condition in which the outcome of infection is largely uniform. Moreover, as a parthenogenetic strain, it reduces experimental variation among individuals due to its uniform genetic background. This reproducibility enables reliable tracking of the parasitism process and facilitates quantitative analyses of host responses and parasitoid-derived factors.

In addition, thelytokous reproduction greatly facilitates genomic analyses by enabling the generation of highly homogeneous genomic DNA, which is critical for high-quality genome assembly⁹. While such parthenogenetic strains are not available in all parasitoid species, our results highlight the potential of Wolbachia-induced thelytoky as a strategy to improve genomic resources. Establishing Wolbachia-infected lines in other parasitoid species may therefore be a powerful approach to expand genomic resources and accelerate comparative molecular studies across parasitoids.

Parasitism outcomes are strongly influenced by host nutritional conditions, indicating that rearing conditions are a critical determinant of parasitoid success. Because successful parasitism depends on precise temporal coordination with host and wasp development, uncontrolled nutritional conditions are likely to disrupt this process and reduce reproducibility. Therefore, careful standardization of host rearing conditions is essential for establishing a reliable experimental system to study parasitoid-host interactions. Furthermore, the fly stock vials should be maintained separately from wasp stock vials, as the presence of wasps reduces fly oviposition behaviors^20,21.

To enable quantitative evaluation of parasitoid-host interactions, we devised a single-oviposition infection assay in which a single egg is deposited into each host. This assay contrasts with conventional bulk infection assays, which often involve multiple oviposition events that lead to excessive host damage and mortality. To promote efficient oviposition within a limited time window (~15 min.), newly-eclosed wasp adults were preconditioned with a glucose and vitamin C solution, thereby improving oviposition efficiency and overall reproducibility.

Furthermore, we optimized the RNAi protocol for gene function analysis in parasitoid wasps. By using the microinjection system, we achieved precise control of injection volume and improved the consistency of knockdown efficiency. Functional validation using IDDF-1 confirmed that this method effectively suppresses gene expression and produces reproducible phenotypic outcomes. Given that transcriptomic and proteomic studies continue to identify large numbers of candidate venom factors, the availability of a functional screening platform will be essential for determining their biological roles.

A limitation of this RNAi protocol is that it is not suitable for analyzing genes that function during the early stages of wasp development, as it is technically challenging to inject dsRNA into wasp eggs that are present inside the host bodies. To overcome this, establishing an ex vivo rearing system may provide a useful approach²². Alternatively, gene knockout approaches using Direct Parental (DIPA)-CRISPR may provide a complementary strategy, whereby reagents are injected into adult females to generate genome-edited offspring²³. Indeed, this approach has successfully generated gene knockout individuals in the pupal ectoparasitoid Nasonia vitripennis by injecting reagents into adult females or individuals at the late pupal stage²⁴. Together with our RNAi-based gene knockdown method, these approaches will facilitate more comprehensive analyses of gene functions involved in the parasitism process.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
50 mL tube	Greiner Bio-One	210261	Infection assay
1 mL syringe	TERUMO	SS-01T	dsRNA injection
Agar	Daisin, Ltd.	P-700	Rearing
Asnol Petri Dish φ40×13.5mm	As One Corporation	1-8549-01	Infection assay
Asnol Petri Dish φ55×17mm	As One Corporation	1-8549-02	dsRNA injection
Blue dye (Erioglaucine disodium salt)	Sigma-Aldrich Co. Llc.	861146-25G	dsRNA injection
Butyl p-Hydroxybenzoate	Nacalai Tesque, Inc.	06327-15	Rearing
CO2 pressure regulator	YAMATOSANGYO	YR-507F-2	Infection assay
Confocal laser scanning microscope	Zeiss	LSM700	Image analysis
Corn meal	Sunny Maize Co., Ltd.	No.4M	Rearing
DAPI	Thermo Fisher Scientific Inc.	PI62247	Nuclei staining, 1:10000
Deionized water	Millipore Inc.	ZLXEV030WW	Infection assay
Dual-Stage Glass Micropipette Puller	Narishige	PC-10	dsRNA injection
Fiji	NA	https://fiji.sc	Image analysis
Filter paper	Advantec	ITEM 526	Rearing
Fly food vial	Chiyoda Science Co., Ltd.	KFB-3M	Rearing
Fly food vial plug	Chiyoda Science Co., Ltd.	AS-275	Rearing
Forceps	Dumont Biologie	11252-20	Dissection
Freezer, -20 °C	Nihon Freezer	GS-3120HC	Sample storage
Freezer, -80 °C	Nihon Freezer	CLN-52UD2	Sample storage
Glass slides	Matsunami Glass Ind., Ltd	S7213	Infection assay
Glucose	Showa Sangyo Co., Ltd.	Not availavle	Rearing
KOD Plus Neo	Toyobo	KOD-401	dsRNA synthesis
Incubator	Panasonic	MIR-254-PJ	Rearing
Ligation high Ver.2	Toyobo	LGK-201	dsRNA synthesis
Micropipette Grinder	Narishige	EG-401	dsRNA injection
Mineral oil	Nacalai Tesque, Inc.	23306-84	dsRNA injection
Nanoject III	Drummond Scientific Company	3-000-207	dsRNA injection
Nanoject Glass Capillaries	Drummond Scientific Company	3-000-203-G/X	dsRNA injection
pBluescript KS (+) plasmid	Not availavle	Not availavle	dsRNA synthesis
pBluescript SK (-) plasmid	Not availavle	Not availavle	dsRNA synthesis
PrimeScript reverse Transcriptase	Takara	2680A	dsRNA synthesis
Propionic acid	Nacalai Tesque, Inc.	29018-55	Rearing
PROWIPE	Daio Paper Corporation	2-2624-02	Infection assay, dsRNA injection.
ReverTra Ace qPCR RT Master Mix with gDNA Remover	Toyobo	FSQ-301	qRT-PCR
RNAiso Plus reagent	Takara	9108	qRT-PCR
Small vial (Test tube 12 mL)	Sarstedt	REF 58.487	Infection assay
Small vial plug	Chiyoda Science Co., ltd.	QD-S4	Infection assay
Spatula	As One Corporation	6-522-02	Rearing
Square petri dish	Eiken Chemical Co., Ltd.	64-2192-55, AW2000	dsRNA injection
Stereomicroscope	Leica Microsystems	lvesta3 (C-Mount)	Infection assay
Stereomicroscope	Nikon Solutions Co., Ltd.	SMZ1000	dsRNA injection
T7 RiboMAX Express RNAi System	Promega	P1700	dsRNA synthesis
Thermal Cycler Dice Real Time System	Takara	TP815	qRT-PCR
Thermal Cycler GeneAtlas	Astel	G02	dsRNA synthesis, qRT-PCR
THUNDERBIRD SYBR qPCR Mix	Toyobo	QPS-201	qRT-PCR
TissueLyser II	Qiagen	Not availavle	qRT-PCR
Vitamin C, L-Ascorbic Acid	Nacalai Tesque, Inc.	03420-65	Rearing
Welch's grape 100 juice	Asahi Soft drinks Co., Ltd.	32390	Infection assay
Yeast	Asahi Group Foods, Ltd.	HB-P02	Rearing