We thank Minoru Moriyama for technical advice and supports, Bin Hirota and Ryutaro Suzuki for collecting Odonata larvae, and Misa Shinya for helpful comments on the manuscript. This work was supported by JSPS KAKENHI Grant Numbers JP18J21561 to GO and JP18H02491, JP18H04893, JP19H03287, and JP20H04936 to RF.

NOTE: The overall scheme of the protocols is shown in Figure 1.
1. Preparation of larvae of dragonflies or damselflies.
<ol>
	<li>Collect larvae in the field using a hand net. 
	NOTE: I. senegalensis larvae often cling onto water plants floating on the water surface, while P. zonata larvae often stay among leaf litter at the bottom. In Tsukuba City, the final instar larvae of I. senegalensis are found mainly from March to June, and those of P. zonata from May to June. I. senegalensis larvae can be reared from eggs in the laboratory14, but the larvae collected in the field have the clearly higher success rate of adult emergence.</li>
	<li>Species identification by restriction fragment length polymorphism (RFLP) of PCR-amplified products. 
	NOTE: This step is only necessary if the species is difficult to identify from the appearance of the larvae, as in I. senegalensis.
	<ol>
		<li>Hold one of the caudal gills of a larva using forceps. Then, the larva falls off its caudal gill itself (causing autotomy). 
		NOTE: Larvae of Zygopteran damselflies usually have three caudal gills (Figure 1A). When they are attacked by a predator, they can take off their own caudal gills to escape. If the caudal gill is not available, a portion of the leg is dissected.</li>
		<li>Put one caudal gill into 100 &#181;L of PBS solution [0.8% NaCl, 0.02% KCl, 0.115% Na2HPO4, and 0.02% KH2PO4 (w/v)] and homogenize with a hand mixer using a deposit pestle.</li>
		<li>Spin down the mixture at 5,000 x g for 10 seconds, and subject 0.5 &#181;L of the supernatant to PCR-amplification using DNA polymerase and primers to amplify the internal transcribed spacer 1 (ITS1) region of the nuclear DNA. 
		NOTE: Perform the PCR according to the manufacturer&#39;s protocol. The combination of ITS-F0 (5&#39;- GGA AAG ATG GCC AAA CTT GA -3&#39;) and 5.8S-AS1 (5&#39;- GCC GGC CCT CAG CCA G -3&#39;) primers can amplify ITS1 region in almost all Odonata species19.</li>
		<li>Incubate the mixture of 1 &#181;L of PCR-amplified product, 0.3 &#181;L of 10x M Buffer, 0.1 &#181;L of DraI restriction enzyme, and 1.6 &#181;L of water at 37 &#176;C for 1 h. 
		NOTE: Select the best restriction enzymes, depending on the combination of species. If there are no restriction enzymes suitable for species identification, confirm by DNA sequencing.</li>
		<li>Load 2 &#181;L of the products with loading dye on 2% agarose gel and run electrophoresis. 
		NOTE: It is difficult to identify species when using 1% or 1.5% agarose gel.</li>
		<li>Check the electrophoresis pattern to identify the species (Figure 2). 
		NOTE: RFLP patterns are species- and population-dependent. In Tsukuba population, I. asiatica has a major band of 400-500 bp, whereas I. senegalensis has a major band of approximately 200 bp and an additional band of less than 100 bp (an arrowhead in Figure 2). The ITS1 region exists in multiple copies in the genome, and in Ischnura species, microsatellite polymorphism within the ITS1 region is often present in the same individual, affecting the pattern of RFLPs.</li>
	</ol>
	</li>
	<li>Rear the collected larvae in the laboratory until use for RNAi.
	<ol>
		<li>Place each larva separately into each well of a 12-well plate with approximately 3 mL of water. 
		NOTE: I. senegalensis larvae must be kept individually because they frequently cannibalize each other, while P. zonata larvae can be kept in a group because they rarely cannibalize.</li>
		<li>Feed I. senegalensis larvae with Artemia brine shrimp every day and P. zonata larvae with bloodworms and/or Tubifex worms at least twice a week until they grow to the suitable developmental stage for RNAi. 
		NOTE: Frequent feeding is critical to increase the success rate of adult emergence.</li>
		<li>Change the water as soon as it becomes dirty with feces or leftovers. 
		NOTE: Frequent water change is also important to increase the success rate of adult emergence.</li>
		<li>Judge the appropriate developmental stage for RNAi. 
		NOTE: The final instar larvae of I. senegalensis can be classified into five developmental stages14,20. The first stage (stage A, before the wings begin to expand) or the second stage (stage B) of the final instar larvae is suitable for RNAi experiments, when considering the clear phenotype of RNAi after adult emergence (see Discussion). In P. zonata, the final instar larvae before wing expansion (corresponding to stage A of I. senegalensis) were used in this study.</li>
	</ol>
	</li>
</ol>
2. Preparation of dsRNA/siRNA solution and injection needles for RNAi.
NOTE: Select either small interfering RNA (siRNA) (step 2.1) or double-stranded RNA (dsRNA) (step 2.2) as the solution for RNAi.
<ol>
	<li>Preparation of the siRNA solution
	<ol>
		<li>Design siRNA using siDirect program version 2.0 (http://sidirect2.rnai.jp/)21,following the guidelines previously reported in Lepidopteran insects22.</li>
		<li>Obtain commercially synthesized siRNA. 
		NOTE: The sequences of siRNA targeting MCO2 gene of I. senegalensis used in this study were as follows: 5&#39;- GCA CUU UCC GUU AUC AAU AUA -3&#39; for sense strand and 5&#39;- UAU UGA UAA CGG AAA GUG CUC -3&#39; for antisense strand. As a negative control, the sequences of siRNA targeting enhanced green fluorescent protein (EGFP) gene were as follows: 5&#39;- GCA UCA AGG UGA ACU UCA AGA -3&#39; for sense strand and 5&#39;- UUG AAG UUC ACC UUG AUG CCG -3&#39; for antisense strand11.</li>
		<li>Dilute the siRNA to 100 &#181;M with injection buffer [100 mM CH3COOK, 2 mM Mg(CH3COO)2, 30 mM HEPES-KOH, pH 7.4]22.</li>
		<li>Store at -80 &#176;C until use.</li>
	</ol>
	</li>
	<li>Preparation of dsRNA solution.
	<ol>
		<li>Select a 300-400 bp region for dsRNA using primer3 program version 4.1.0 (http://bioinfo.ut.ee/primer3/)23 and design the primer sets.&#160;</li>
		<li>Obtain commercially synthesized primer sets. 
		&#8203;NOTE: To produce templates for dsRNA synthesis, the primer sets used in this study were as follows: (5&#39;- GCC TGT CAG CTT TGT CTT CC -3&#39; for forward primer and 5&#39;- GGT GTC TGG CGG ACA ACT AT -3&#39; for reverse primer) for MCO2 genes of I. senegalensis (IsMCO2, accession no. LC589180) and (5&#39;- CCG CAC AGC TCA CTA TTC AA -3&#39; for forward primer and 5&#39;- GGA GGA TTC CTT CAT CGA CA -3&#39; for reverse primer) for MCO2 genes of P. zonata (PzMCO2, accession no. LC589179). As a negative control, the primer set for dsRNA targeting &#946;-lactamase (bla) gene on the cloning vector were (5&#39;- CTA TGT GGC GCG GTA TTA T -3&#39; for forward primer and 5&#39;- CAG AAG TGG TCC TGC AAC T -3&#39; for reverse primer).</li>
		<li>Extract RNA from Odonata larvae using a commercially available RNA extraction kit and perform cDNA synthesis using reverse transcriptase according to the manufacturer&#39;s protocol. 
		NOTE: cDNA library prepared for RNA sequencing analysis can be also used.</li>
		<li>Amplify the target sequences using the synthesized cDNA and the designed primer set and clone them into the cloning vector using a commercial ligase according to the manufacturer&#39;s protocol.</li>
		<li>Transform the plasmid into E. coli competent cells and pick up a single colony after overnight incubation.</li>
		<li>PCR-amplify the insert region using primers on the vector.</li>
		<li>Confirm the cloned sequence by Sanger sequencing.</li>
		<li>PCR-amplify the insert using vector primers containing the T7 polymerase promoter sequence24, 25. 
		NOTE: In this study, the following primers were used: T7-F (5&#39; - TAA TAC GAC TCA CTA TAG GGA GAC TAG TCA TAT GGA T - 3&#39;) and T7-R (5&#39;- TAA TAC GAC TCA CTA TAG GGA GAC CCG GGG ATC CGA T - 3&#39;) 25.</li>
		<li>Purify the PCR product using the PCR purification kit according to the manufacturer&#39;s protocol, elute the DNA with 50 &#181;L of distilled water, and concentrate the eluted DNA solution to approximately 10 &#181;L using a centrifugal evaporator.</li>
		<li>Synthesize dsRNA by in vitro transcription according to the manufacturer&#39;s protocol. Use a total of 1000 ng template DNA and elute synthesized dsRNA with 100 &#181;L of elution buffer.</li>
		<li>Measure the concentration of dsRNA using a spectrophotometer and confirm the quality of dsRNA by electrophoresis on a 1.5% agarose gel. 
		NOTE: A single band can be seen when dsRNA synthesis is successful.</li>
		<li>Dilute dsRNA to 1000 ng/&#181;L with elution buffer and store at -20 &#176;C until use.</li>
	</ol>
	</li>
	<li>Preparation of injection needles
	<ol>
		<li>Pull a glass capillary by using a glass needle puller.</li>
		<li>Place the tip of the pulled capillary onto a double-sided adhesive tape and break the tip of the capillary with forceps.</li>
		<li>Set the capillary to an injector. 
		NOTE: It is easier to handle the capillary if you wear nitrile gloves.</li>
		<li>Load siRNA/dsRNA solution to the prepared capillary. 
		NOTE: Do not use the capillary repeatedly since the tip of the injected capillary may become clogged with dirt because the larvae collected in the field lived in the mud.</li>
	</ol>
	</li>
</ol>
3. siRNA/dsRNA injection
NOTE: The procedure is slightly different for damselflies (step 3.1) and dragonflies (step 3.2).
<ol>
	<li>Injection to a Zygopteran (damselfly) larva (e.g., I. senegalensis).
	<ol>
		<li>Anesthetize a larva covered with a wet paper on crushed ice for 50-70 seconds (Figure 3A-C). 
		NOTE: Duration of ice-cold anesthesia depends on the condition of the larva; if the larva begins to move after 70 seconds of ice-cold anesthesia, an additional 70 seconds of ice-cold anesthesia is applied. For larvae that rarely move (e.g., P. zonata), this procedure is not essential.</li>
		<li>Attach two pins on both sides of the prothorax and fix the larva to a fixed stand (e.g., a piece of Styrofoam) (Figure 3D). 
		NOTE: Adjust the position and number of pins, depending on the place of injection.</li>
		<li>Stretch the inter-segmental membrane between the 7th and 8th abdominal segment for RNAi in the abdomen or between the prothorax and synthorax (fused mesothorax and metathorax) for RNAi in the thorax using hands and forceps.</li>
		<li>Keep the inter-segmental membrane stretched by hand.</li>
		<li>Insert the tip of the prepared capillary into the stretched inter-segmental membrane (Figure 3D, 3F).</li>
		<li>Inject 1 &#181;L of siRNA/dsRNA solution.</li>
	</ol>
	</li>
	<li>Injection to an Anisopteran (dragonfly) larva (e.g., P. zonata).
	<ol>
		<li>Wipe the water off the larval surface with a paper towel.</li>
		<li>If needed, attach two pins on both sides of the prothorax and fix the larva to a fixed stand (e.g., a piece of Styrofoam) (Figure 3H). 
		NOTE: Adjust the position and number of pins, depending on the place of injection. In the case of P. zonata, it is not essential to fix the larvae with pins at this step.</li>
		<li>Stretch the inter-segmental membrane between the 4th and 5th abdominal segment by hand.</li>
		<li>Make a small hole with a fine needle in the inter-segmental membrane between the 4th and 5th abdominal segment. 
		NOTE: This procedure is necessary for many Anisopteran (dragonfly) species because the inter-segmental membrane is too hard to insert a glass capillary directly.</li>
		<li>Insert the tip of the prepared capillary into the prepared hole (Figure 3I). 
		NOTE: As shown in Figure 3H, part of the dorsal side of the larvae corresponds to the ventral side of the adult, so the phenotype appears ventrally when treated as shown in Figure 3H-J.</li>
		<li>Inject 1 &#181;L of siRNA/dsRNA solution.</li>
	</ol>
	</li>
</ol>
4. in vivo electroporation
<ol>
	<li>If needed, add more pins to fix the larva to a fixed stand (e.g., a piece of Styrofoam).</li>
	<li>Following the injection of the siRNA/dsRNA solution, apply two droplets of ultrasound gel on the larval surface using forceps.</li>
	<li>Place electrodes on the ultrasound gel, with the positive electrode on the side injected with the siRNA/dsRNA solution and a negative electrode on the opposite side (Figure 3E, 3G, 3J). 
	NOTE: Do not directly touch the epidermis of the larva to avoid the burning effect of electroporation.</li>
	<li>Generate 10-times electroporation pulses (280 ms/s each) using an electroporator. 
	NOTE: Adjust the voltage of electroporation, depending on the species, stages and tissues. In this study, 25 V was applied to I. senegalensis and 45 V to P. zonata.</li>
	<li>Wipe off the remaining gel on the surface with a paper towel.</li>
	<li>Keep the treated larvae resting on a wet paper towel for approximately one day for recovery and transfer them to a rearing case on the following day.</li>
</ol>
5. Site-specific phenotypic analysis
<ol>
	<li>Keep I. senegalensis individually in a Petri dish (5 cm in diameter) containing approximately 10 mL of water and a piece of paper towel, and keep P. zonata in a plastic cage with a disposable non-woven mesh (eclosion cage14).</li>
	<li>For I. senegalensis, after the larvae stop eating, move them individually into a plastic cage with a disposable non-woven mesh. 
	NOTE: Do not put two or more larvae in the same cage. Otherwise, they will cannibalize each other immediately after they become adults.</li>
	<li>After the emergence of the adult, observe and photograph the phenotype around the region where the positive electrode was placed for electroporation. 
	NOTE: The phenotype appears only in patches. Phenotypes are often difficult to recognize immediately after emergence due to incomplete pigmentation.</li>
	<li>In order to examine the efficiency of RNAi (e.g., quantitative RT-PCR), if the phenotype is visible, dissect the region and compare it with the region without the phenotype. 
	NOTE: The levels of RNAi phenotype, namely size and location of the cuticle decolorization, often exhibit considerable variation between individuals (see Figure 4).</li>
	<li>Keep the emerged adults in 100% EtOH for future analyses. 
	NOTE: Body color of Odonata species quickly fades after death, so it is important to store them in ethanol before they die. Ethanol sometimes discolors the insects, in such cases that the insects are frozen before they die.</li>
</ol>

<ol>
	<li>Misof, B., 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=Phylogenomics+resolves+the+timing+and+pattern+of+insect+evolution.">Phylogenomics resolves the timing and pattern of insect evolution.</a> Science. 346 (6210), 763-767 (2014).</li><li>Wipfler, B., 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=Evolutionary+history+of+Polyneoptera+and+its+implications+for+our+understanding+of+early+winged+insects.">Evolutionary history of Polyneoptera and its implications for our understanding of early winged insects.</a> Proceedings of the National Academy of Sciences of the United States of America. 116 (8), 3024-3029 (2019).</li><li>Corbet, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Dragonflies Behavior and Ecology of Odonata. , (1999).</li><li>Futahashi, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Color+vision+and+color+formation+in+dragonflies.">Color vision and color formation in dragonflies.</a> Current Opinion in Insect Science. 17, 32-39 (2016).</li><li>Futahashi, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diversity+of+UV+reflection+patterns+in+Odonata.">Diversity of UV reflection patterns in Odonata.</a> Frontiers in Ecology and Evolution. 8 (201), (2020).</li><li>C&#243;rdoba-Aguilar, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Dragonflies and damselflies. Model organisms for ecological and evolutionary research. , (2008).</li><li>Bybee, S., 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=Odonata+(dragonflies+and+damselflies)+as+a+bridge+between+ecology+and+evolutionary+genomics.">Odonata (dragonflies and damselflies) as a bridge between ecology and evolutionary genomics.</a> Frontiers in Zoology. 13 (46), (2016).</li><li>Linz, D. M., Clark-Hachtel, C. M., Borr&#224;s-Castelles, F., Tomoyasu, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Larval+RNA+interference+the+red+flour+beetle,+Tribolium+castaneum.">Larval RNA interference the red flour beetle, Tribolium castaneum.</a> Journal of Visualized Experiments. 92, 52059 (2014).</li><li>Okude, G., 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=Electroporation-mediated+RNA+interference+reveals+a+role+of+multicopper+oxidase+2+gene+in+dragonfly&#8217;s+cuticular+pigmentation.">Electroporation-mediated RNA interference reveals a role of multicopper oxidase 2 gene in dragonfly&#8217;s cuticular pigmentation.</a> Applied Entomology and Zoology. 53 (3), 379-387 (2017).</li><li>Terenius, O., 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=RNA+interference+in+Lepidoptera:+An+overview+of+successful+and+unsuccessful+studies+and+implications+for+experimental+design.">RNA interference in Lepidoptera: An overview of successful and unsuccessful studies and implications for experimental design.</a> Journal of Insect Physiology. 57, 231-245 (2011).</li><li>Ando, T., Fujiwara, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroporation-mediated+somatic+transgenesis+for+rapid+functional+analysis+in+insects.">Electroporation-mediated somatic transgenesis for rapid functional analysis in insects.</a> Development. 140, 454-458 (2013).</li><li>Nishikawa, H., 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=A+genetic+mechanism+for+female-limited+Batesian+mimicry+in+Papilio+butterfly.">A genetic mechanism for female-limited Batesian mimicry in Papilio butterfly.</a> Nature Genetics. 47 (4), 405-409 (2015).</li><li>Osanai-Futahashi, M., 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=Positional+cloning+of+a+Bombyx+pink-eyed+white+egg+locus+reveals+the+major+role+of+cardinal+in+ommochrome+synthesis.">Positional cloning of a Bombyx pink-eyed white egg locus reveals the major role of cardinal in ommochrome synthesis.</a> Heredity. 116 (2), 135-145 (2016).</li><li>Okude, G., Futahashi, R., Tanahashi, M., Fukatsu, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laboratory+rearing+system+for+Ischnura+senegalensis+(Insecta:+Odonata)+enables+detailed+description+of+dragonfly&#8217;s+larval+development+and+morphogenesis.">Laboratory rearing system for Ischnura senegalensis (Insecta: Odonata) enables detailed description of dragonfly&#8217;s larval development and morphogenesis.</a> Zoological Science. 34 (5), 386-397 (2017).</li><li>Ozono, A., Kawashima, I., Futahashi, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Dragonflies of Japan (3rd edition). , (2017).</li><li>Ozono, A., Kawashima, I., Futahashi, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Handbook of Japanese Aquatic Insects. Volume 3: Dragonfly larvae. , (2019).</li><li>Arakane, Y., Noh, M. Y., Asano, T., Kramer, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tyrosine+metabolism+for+insect+cuticle+pigmentation+and+sclerotization.">Tyrosine metabolism for insect cuticle pigmentation and sclerotization.</a> Extracellular Composite Matrices in Arthropods. , 165-220 (2016).</li><li>Asano, 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=Mini-review+an+insect-specific+system+for+terrestrialization:+Laccase-mediated+cuticle+formation.">Mini-review an insect-specific system for terrestrialization: Laccase-mediated cuticle formation.</a> Insect Biochemistry and Molecular Biology. 108, 61-70 (2019).</li><li>Futahashi, R., Okude, G., Sugimura, M., Ugai, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interspecific+hybrids+in+Japanese+Odonata.">Interspecific hybrids in Japanese Odonata.</a> Tombo. 60, 1-49 (2018).</li><li>Okude, G., Fukatsu, T., Futahashi, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interspecific+crossing+between+blue-tailed+damselflies+Ischnura+elegans+and+I.+senegalensis+in+the+laboratory.">Interspecific crossing between blue-tailed damselflies Ischnura elegans and I. senegalensis in the laboratory.</a> Entomological Science. 23 (2), 165-172 (2020).</li><li>Naito, Y., Yoshimura, J., Morishita, S., Ui-Tei, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=siDirect+2.0:+updated+software+for+designing+functional+siRNA+with+reduced+seed-dependent+off-target+effect.">siDirect 2.0: updated software for designing functional siRNA with reduced seed-dependent off-target effect.</a> BMC Bioinformatics. 10, 392 (2009).</li><li>Yamaguchi, J., Mizoguchi, T., Fujiwara, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=siRNAs+induce+efficient+RNAi+response+in+Bombyx+mori+embryos.">siRNAs induce efficient RNAi response in Bombyx mori embryos.</a> PLoS ONE. 6 (9), e25469 (2011).</li><li>Untergasser, A., 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=Primer3--new+capabilities+and+interfaces.">Primer3--new capabilities and interfaces.</a> Nucleic Acids Research. 40 (15), e115 (2012).</li><li>Futahashi, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole-mount+in+situ+hybridization+of+sectioned+tissues+of+species+hybrids+to+detect+cis-regulatory+changes+in+gene+expression+pattern.">Whole-mount in situ hybridization of sectioned tissues of species hybrids to detect cis-regulatory changes in gene expression pattern.</a> Methods in Molecular Biology. 772, 319-328 (2011).</li><li>Matsuura, Y., Kikuchi, Y., Miura, T., Fukatsu, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrabithorax+is+essential+for+bacteriocyte+development.">Ultrabithorax is essential for bacteriocyte development.</a> Proceedings of the National Academy of Sciences of the United States of America. 112 (30), 9376-9381 (2015).</li><li>Henze, M. J., Lind, O., Wilts, B. D., Kelber, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pterin-pigmented+nanospheres+create+the+colours+of+the+polymorphiic+damselfly+Ischnura+elegans.">Pterin-pigmented nanospheres create the colours of the polymorphiic damselfly Ischnura elegans.</a> Journal of The Royal Society Interface. 16, 20180785 (2019).</li><li>Nishide, Y., 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=Diversity+and+function+of+multicopper+oxidase+genes+in+the+stinkbug+Plautia+stali.">Diversity and function of multicopper oxidase genes in the stinkbug Plautia stali.</a> Scientific Reports. 10 (1), 3464 (2020).</li><li>Christiaens, O., Swevers, L., Smagghe, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DsRNA+degradation+in+the+pea+aphid+(Acyrthosiphon+pisum)+associated+with+lack+of+response+in+RNAi+feeding+and+injection+assay.">DsRNA degradation in the pea aphid (Acyrthosiphon pisum) associated with lack of response in RNAi feeding and injection assay.</a> Peptides. 53, 307-314 (2014).</li><li>Sun, D., Guo, Z., Liu, Y., Zhang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Progress+and+prospects+of+CRISPR/Cas+systems+in+insects+and+other+arthropods.">Progress and prospects of CRISPR/Cas systems in insects and other arthropods.</a> Frontiers in Physiology. 8, 608 (2017).</li><li>Miyakawa, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+study+of+the+life-history+of+Pseudothemis+zonata+(Burm.)+(Odonata,+Libellulidae)+II.+Immature+stage.">A study of the life-history of Pseudothemis zonata (Burm.) (Odonata, Libellulidae) II. Immature stage.</a> Konty&#251;. 37 (4), 409-422 (1969).</li></ol>

The authors have nothing to disclose.

Lethality of RNAi treatment
We found that the lethality of the RNAi treatment depends strongly on the rearing history and condition of the Odonata larvae. The larvae soon after collection in the field are generally healthy and exhibit low mortality rates after the electroporation-mediated RNAi treatment. By contrast, the larvae reared in the laboratory for a long period of time (e.g., one month) tend to suffer low success rates of adult emergence. In I. senegalensis, instead of the larvae collected in the field, the larvae reared in the laboratory from eggs can be used14, but the success rates of RNAi using the laboratory-reared larvae tend to be considerably lower (many individuals died during metamorphosis) than those using the field-collected larvae. In addition, frequent larval feeding and clean water rearing are important for increasing the success rates of adult emergence and reducing the lethality of the RNAi treatment.
Efficiency of RNAi treatment
As described above, the levels of RNAi phenotype, namely size and location of the cuticle decolorization, often exhibited considerable variation between individuals subjected to the same RNAi treatment (e.g., Figure 4), but the levels of the phenotypic penetrance seem to be remarkably different between the Odonata species. The observed phenotypic regions were larger and more prominent in I. senegalensis (Figures 4-5) than in P. zonata (Figure 6) and N. pygmaea9. This difference may be due to the thickness of the cuticle on the larval surface, considering that the cuticle of I. senegalensis is thinner than the cuticle of P. zonata and N. pygmaea). As far as we examined, no clear difference was recognized between the effects of siRNA and dsRNA (Figure 4, Table 1).
Appropriate developmental stage for RNAi
It should be noted that proper larval staging is important for performing RNAi efficiently. Inhibition of adult pigmentation was caused by MCO2 RNAi before the stage D (approximately 3 days before adult emergence), which is consistent with the previous report on N. pygmaea9. The RNAi phenotypes observed when injected at the stages C and D were less conspicuous than those treated at the stages A and B, which indicate that the stages C and D may be too late to sufficiently suppress the gene expression. The appropriate timing for RNAi treatment depends on the timing of gene expression, and MCO2 gene exhibits transiently high expression during adult emergence9, as in other insects17,18. In the stinkbug Plautia stali, RNAi knockdown of MCO2 gene was observed from day 4 onwards after injection27, which is consistent with the present results.
Our previous study on I. senegalensis showed that, after the stage B, days to adult emergence exhibit relatively small variation among the majority of final instar larvae, suggesting that the stage B may correspond to the onset of the process toward adult emergence, after which the developmental processes for metamorphosis proceed in a prefixed and coordinated manner14. Morphological abnormalities caused by wounds were often observed when the larvae were RNAi-treated at the stages C and D (Figure 7B, 7C). This is likely to be associated with a dramatic progression of metamorphosis during these stages, suggesting that RNAi treatment should be avoided from the stage C and on. In summary, we recommend that final instar larvae at the stage A or B (or at the stage before the larval wings expand significantly) should be used for RNAi experiments.
Usefulness and superiority of electroporation-mediated RNAi method
The conventional RNAi is a simple and powerful experimental method, but some insect lineages like butterflies10, aphids28 and dragonflies9 exhibit low RNAi efficiency, for which establishment of gene function analysis is a major challenge. In this study, we found that electroporation-mediated RNAi can induce local gene suppression in dragonflies with almost 100% efficiency, at least in epidermis, if treated at appropriate developmental stages (Table 1). Recently, CRISPR/Cas9-based gene knockouts have been successfully applied to a variety of insects, providing a powerful molecular genetic tool for non-model organisms29. Here, however, we point out that CRISPR/Cas9 is certainly great but the electroporation-mediated RNAi method may be superior to CRISPR/Cas9 in some respects.
Firstly, in the electroporation-mediated RNAi method, the body region where RNAi phenotypes appear can be easily controlled experimentally by the position of the positive electrode upon electroporation. In addition, since the region where the gene expression is suppressed is limited around the region where the positive electrode was placed, the RNAi phenotypes can be easily compared with the control phenotypes side by side in the same individual. Secondly, compared to CRISPR/Cas9 method in which injected eggs have to be reared to adulthood to observe the knockout phenotypes, the electroporation-mediated RNAi is superior in that the gene knockdown phenotypes can usually be observed in much shorter time. For example, it takes three to four months for I. senegalensis and one to two years for P. zonata from eggs to adults14,30. However, in order to observe RNAi phenotypes within the adult epidermis, it takes less than one month from dsRNA injection into final instar larvae at the stage B to adult emergence for both I. senegalensis and P. zonata (Figure 7). Thirdly, the electroporation-mediated RNAi method entails dsRNA injection into large larvae, which is easier than microinjection into tiny eggs required for CRISPR/Cas9 method. In addition, the electroporation-mediated RNAi is applicable to insect species whose newly laid eggs are difficult to collect. For example, females of P. zonata lay eggs onto floating plants on water surface during flight, and thus it is difficult to collect their eggs both in the field and in the lab. Hence, we expect that this protocol may be generally applicable to non-model organisms in which the conventional RNAi method does not work efficiently.

Dragonflies and damselflies (the order Odonata) are among the most ancestral groups of insects that exhibit &#34;metamorphosis&#34;1,2. By metamorphosis, they change their habitat, morphology, and behavior drastically from aquatic larvae to terrestrial/aerial adults3. Odonata adults have a well-developed color vision and represent a remarkable diversity in body colors and patterns across sexes, stages, and species3,4,5. While many ecological and behavioral studies on Odonata have been conducted6,7, molecular genetic studies have been hindered mainly by the difficulty in applying gene functional analysis to Odonata.
The conventional RNA interference (RNAi) method, in which double-stranded RNA (dsRNA) is injected to suppress the function of the gene of interest8, turned out to be ineffective in Odonata insects9, as reported in Lepidopteran insects10. On the other hand, previous reports have suggested that electroporation-mediated RNAi is effective in Lepidopteran species, especially in epidermal tissues11,12,13. We recently found that the electroporation-mediated RNAi works effectively in the tiny dragonfly Nannophya pygmaea (Libellulidae: Anisoptera)9, but N. pygmaea is a relatively rare species and therefore not suitable for molecular genetic studies.
Most Odonata species are classified into either of the two suborders, Zygoptera (damselflies) or Anisoptera (true dragonflies)3. Here we focused on the blue-tailed damselfly Ischnura senegalensis (Coenagrionidae; Figure 1A) as a representative Zygopteran species and the pied skimmer dragonfly Pseudothemis zonata (Libellulidae; Figure 1B) as a representative Anisopteran species. The two species are among the most common Odonata species in natural and urban ponds in Japan, including those in Tsukuba City, and we can collect many larvae of the two species in the field. Recently we established a laboratory rearing system for individual larvae of I. senegalensis, which enabled continuous monitoring of development and morphogenesis of the Odonata larvae in detail14.
In this report, we provide a refined method and a video protocol for the electroporation-mediated RNAi in I. senegalensis and P. zonata. In Japan, I. senegalensis and Ischnura asiatica, which are genetically close, are often found sympatrically15, and they are difficult to distinguish in larvae16. We also describe how two Ischnura species can be distinguished by restriction fragment length polymorphism (RFLP).
For evaluating the effectiveness of the electroporation-mediated RNAi, we select multicopper oxidase 2 gene (MCO2; also known as laccase2) as a representative target gene, on account of the visible phenotype of paler cuticle color upon knockdown of the gene expression9. MCO2 is known to be essential for darkening of the epidermis in a variety of insect species17,18.

<table><tbody><tr><td>12-well plate</td><td>Violamo</td><td>VTC-P12</td><td>For rearing larvae before injection</td></tr><tr><td>Brine shrimp eggs</td><td>JAPAN PET DESIGN</td><td>4975677012396</td><td></td></tr><tr><td>Calibrated micropipette (1-5 &amp;micro;L)</td><td>Drummond</td><td>2-000-001-90</td><td></td></tr><tr><td>Deposit pestle 1.5 mL</td><td>Thermo Fisher Scientific</td><td>K749520-0090</td><td></td></tr><tr><td>Digital high defenition microscope camera</td><td>Leica Microsystems</td><td>MC170HD</td><td></td></tr><tr><td>Disposable non-woven mesh</td><td>HAKUGEN EARTH</td><td>DSC-105A</td><td></td></tr><tr><td>DNA Ligation Kit Ver.2.1</td><td>Takara Bio</td><td>6022</td><td></td></tr><tr><td>DraI</td><td>Takara Bio</td><td>1037A</td><td></td></tr><tr><td>Electrode (1mm&amp;phi;)</td><td>NEPAGENE</td><td>CUY650P1</td><td></td></tr><tr><td>Electroporator</td><td>CellProduce</td><td>Cure-gene</td><td></td></tr><tr><td>Forceps</td><td>KOWA Forceps&amp;nbsp;Industry</td><td>K-10 No1</td><td></td></tr><tr><td>Glass needle puller</td><td>NARISHIGE</td><td>PN-3</td><td></td></tr><tr><td>Hand mixer</td><td>AS ONE</td><td>1-229-02</td><td></td></tr><tr><td>Hand net</td><td>HOGA</td><td>IS33-3B</td><td></td></tr><tr><td>HEPES</td><td>FUJIFILM Wako&amp;nbsp;Pure Chemical Corporation</td><td>346-01373</td><td>For injection buffer</td></tr><tr><td>Insect pin capitate No. 3</td><td>Shiga Konchu Fukyusha</td><td>N230</td><td>For fixing larvae</td></tr><tr><td>KCl</td><td>FUJIFILM Wako&amp;nbsp;Pure Chemical Corporation</td><td>163-03545</td><td>For PBS</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>FUJIFILM Wako&amp;nbsp;Pure Chemical Corporation</td><td>169-04245</td><td>For PBS</td></tr><tr><td>KOAc</td><td>FUJIFILM Wako&amp;nbsp;Pure Chemical Corporation</td><td>160-03175</td><td>For injection buffer</td></tr><tr><td>KOH</td><td>FUJIFILM Wako&amp;nbsp;Pure Chemical Corporation</td><td>168-21815</td><td>For injection buffer</td></tr><tr><td>Laboratory Jack (150x150)</td><td>AS ONE</td><td>1-4642-11</td><td>For adjusting the position of the manipulator</td></tr><tr><td>LOGIQLEAN Gel for Ultrasound Hard type</td><td>GE Healthcare</td><td>2369385</td><td></td></tr><tr><td>Manipulator</td><td>Muromachi Kikai Co., Ltd.</td><td>SJ-1</td><td>For adjusting the position of the injector and the capillary</td></tr><tr><td>MEGAscript RNAi kit</td><td>Thermo Fisher Scientific</td><td>AM1626</td><td></td></tr><tr><td>Mg(OAc)&lt;sub&gt;2&lt;/sub&gt;</td><td>FUJIFILM Wako&amp;nbsp;Pure Chemical Corporation</td><td>130-00095</td><td>For injection buffer</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>FUJIFILM Wako&amp;nbsp;Pure Chemical Corporation</td><td>197-02865</td><td>For PBS</td></tr><tr><td>NaCl</td><td>FUJIFILM Wako&amp;nbsp;Pure Chemical Corporation</td><td>191-01665</td><td>For PBS</td></tr><tr><td>Petri dish (5 cm diameter)</td><td>Iwaki</td><td>1010-060</td><td>For rearing injected larvae</td></tr><tr><td>Pneumatic Injector</td><td>NARISHIGE</td><td>IM-12</td><td></td></tr><tr><td>pT7Blue T-Vector</td><td>Novagen</td><td>69820</td><td></td></tr><tr><td>QIAquick PCR Purification Kit</td><td>QIAGEN</td><td>28106</td><td></td></tr><tr><td>RNAiso Plus</td><td>FUJIFILM Wako&amp;nbsp;Pure Chemical Corporation</td><td>9109</td><td></td></tr><tr><td>RNeasy Mini Kit</td><td>QIAGEN</td><td>74106</td><td></td></tr><tr><td>Shiga micro insect pin with stainless headless</td><td>Shiga Konchu Fukyusha</td><td>N251</td><td>For making a small hole</td></tr><tr><td>Stereoscopic microscope</td><td>Leica Microsystems</td><td>S8APO</td><td></td></tr><tr><td>SuperScript IV Reverse Transcriptase</td><td>Thermo Fisher Scientific</td><td>18090010</td><td></td></tr><tr><td>TaKaRa Ex Taq</td><td>Takara Bio</td><td>RR001B</td><td>For PCR-amplification from plasmid</td></tr><tr><td>Tks Gflex DNA Polymerase</td><td>Takara Bio</td><td>R060B</td><td>For PCR-amplification from caudal gill</td></tr></tbody></table>

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 applied the above protocol to electroporation-mediated RNAi targeting MCO2 gene and negative control genes (EGFP for siRNA and bla for dsRNA) (i) in the abdomen of I. senegalensis (Figure 4), (ii) in the thorax of I. senegalensis (Figure 5), and (iii) in the abdomen of P. zonata (Figure 6). The results of the RNAi experiments are summarized in Table 1. Because negatively charged siRNA/dsRNA is incorporated only into positively charged cells, RNAi phenotypes were observed around the region where the positive electrode was placed for electroporation.
In both I. senegalensis and P. zonata, inhibition of melanin pigmentation (i.e., black, brown, and reddish brown) appeared in patches around the region where the positive electrode was placed (white arrowheads and dotted lines in Figure&#160;4, Figure 5,&#160;and Figure 6) when MCO2 RNAi was performed in combination with electroporation (Table 1), as previously reported in N. pygmaea9. By contrast, no phenotypic effects were observed around the electroporation site when the control gene was injected (EGFP siRNA or bla dsRNA) (Figure&#160;4,&#160;Figure 5,&#160;and&#160;Figure 6, Table 1). In addition, injecting the MCO2 gene without electroporation had no effect on adult pigmentation (Figure 4, Table 1), indicating that electroporation is essential for RNAi in Odonata. It should be noted that the blue, green, and yellow colorations are not affected by the RNAi of MCO2 gene that is involved in melanin synthesis in the cuticle, which plausibly reflect the fact that these body colors are attributed to pigment granules present in the epidermal cells that are visible through the transparent cuticle26. As shown in Figure 4, no remarkable phenotypic differences were recognized between the individuals subjected to siRNA treatment and dsRNA treatment, whereas considerable variation in size and location of the RNAi phenotype was observed among different individuals subjected to the same RNAi treatment (ex. compare two examples of IsMCO2 dsRNA in Figure 4).
To determine the developmental stage most suitable for the RNAi treatment, we compared the phenotypic consequences of the RNAi treatment at five morphological stages (stage A-E) in the final larval instar of I. senegalensis (Figure 7A). Inhibition of melanin pigmentation caused by MCO2 RNAi was observed in all emerged adults when injected at the stages A and B (Figure 7C). When injected at the stages C and D, suppression of melanin pigmentation was certainly observed in some emerged adults, but other adults exhibited abnormal coloration caused by wounds (Figure 7B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61952/61952fig1.jpg" /> 
Figure 1: Electroporation-mediated RNAi methods in Odonata. A.&#160;Ischnura senegalensis (Coenagirionidae) as a representative damselfly species. B. Pseudothemis zonata (Libellulidae) as a representative dragonfly species. C. The overall scheme of the protocols. Blue and orange boxes indicate the protocols for I. senegalensis and P. zonata, respectively. The purple boxes indicate the common protocols applied to both species. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61952/61952fig1large.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/61952/61952fig2.jpg" /> 
Figure 2: A representative result of restriction fragment length polymorphism (RFLP)-based species identification for Ischnura species. Arrowhead indicates the I. senegalensis-specific band. 1, 2, 4: I. asiatica, 3: I. senegalensis, M: 100-base pair ladder marker. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61952/61952fig2large.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/61952/61952fig3.jpg" /> 
Figure 3: Electroporation-mediated RNAi method in Odonata. A-C. Ice-cold anesthesia of I. senegalensis. Arrowheads indicate a larva. A. Putting a larva on crushed ice with a wet paper. B. Magnified view of a larva on ice. C. A larva covered with a wet paper on ice. D-G. RNAi method for I. senegalensis. D. Injection into the thorax. E. Electroporation on the thorax. F. Injection into the abdomen. G. Electroporation on the abdomen. H-J. RNAi method for P. zonata. H. Making a small hole on the abdomen. I. Injection into the abdomen. J. Electroporation on the abdomen. Arrows indicate the point of making a hole or injection. +, -: Positive/negative side electrodes. Numbers indicate the abdominal segment. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61952/61952fig3large.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/61952/61952fig4.jpg" /> 
Figure 4: Dorsal views of RNAi phenotypes on the abdomen of I. senegalensis. White arrowheads indicate the regions of suppressed melanization. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61952/61952fig4large.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/61952/61952fig5.jpg" /> 
Figure 5: Lateral and dorsal views of RNAi phenotypes on the thorax of I. senegalensis. White arrowheads and dotted lines indicate the regions of suppressed pigmentation. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61952/61952fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61952/61952fig6.jpg" /> 
Figure 6: Ventral views of RNAi phenotypes in the abdomen of P. zonata. White arrowheads and dotted lines indicate the regions of suppressed melanization. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61952/61952fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61952/61952fig7.jpg" /> 
Figure 7: Stage dependent IsMCO2 RNAi effects during the final larval instar of I.senegalensis.&#160;A.&#160;Morphological changes in the compound eyes at five morphological stages (stage A-E) and the number of days to adult emergence in this study. Numbers in parentheses are from previous report14. B.&#160;Abnormal pigmentation due to wounds. Arrowhead indicates electroporation site. C.&#160;The effect of RNAi at five morphological stages on adult pigmentation in I. senegalensis. The number on the bar indicates the number of individuals.&#160;<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61952/61952fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Species</td>
			<td colspan="7">I.senegalensis</td>
			<td colspan="3">P.zonata</td>
		</tr>
		<tr>
			<td colspan="2">Injected region</td>
			<td colspan="5">Abdomen</td>
			<td colspan="2">Thorax</td>
			<td colspan="3">Abdomen</td>
		</tr>
		<tr>
			<td colspan="2">siRNA/dsRNA</td>
			<td colspan="2">siRNA</td>
			<td colspan="3">dsRNA</td>
			<td colspan="2">dsRNA</td>
			<td colspan="3">dsRNA</td>
		</tr>
		<tr>
			<td colspan="2">Target gene</td>
			<td>IsMCO2</td>
			<td>EGFP</td>
			<td>IsMCO2</td>
			<td>IsMCO2</td>
			<td>bla</td>
			<td>IsMCO2</td>
			<td>bla</td>
			<td>PzMCO2</td>
			<td>PzMCO2</td>
			<td>bla</td>
		</tr>
		<tr>
			<td colspan="2">Electroporation</td>
			<td>+</td>
			<td>+</td>
			<td>+</td>
			<td>-</td>
			<td>+</td>
			<td>+</td>
			<td>+</td>
			<td>+</td>
			<td>-</td>
			<td>+</td>
		</tr>
		<tr>
			<td colspan="2">Injected larvae</td>
			<td>22</td>
			<td>25</td>
			<td>30</td>
			<td>6</td>
			<td>53</td>
			<td>12</td>
			<td>20</td>
			<td>17</td>
			<td>5</td>
			<td>9</td>
		</tr>
		<tr>
			<td colspan="2">Emerged adults</td>
			<td>7</td>
			<td>6</td>
			<td>13</td>
			<td>4</td>
			<td>40</td>
			<td>11</td>
			<td>14</td>
			<td>11</td>
			<td>2</td>
			<td>5</td>
		</tr>
		<tr>
			<td colspan="2">Adults with less pigmented regions (ratio)</td>
			<td>7 
			(100%)</td>
			<td>0 
			(0%)</td>
			<td>13 
			(100%)</td>
			<td>0 
			(0%)</td>
			<td>0 
			(0%)</td>
			<td>10 
			(91%)</td>
			<td>0 
			(0%)</td>
			<td>11 
			(100%)</td>
			<td>0 
			(0%)</td>
			<td>0 
			(0%)</td>
		</tr>
	</tbody>
</table>
Table 1. The effect of RNAi on adult pigmentation in I. senegalensis and P. zonata. Results at stage A are shown in I. senegalensis. IsMCO2: multicopper oxidase 2 gene of I. senegalensis, EGFP: Enhanced green fluorescent protein gene, bla: beta lactamase gene from pGEM-T Easy Vector, PzMCO2: multicopper oxidase 2 gene of P. zonata. The results for the control genes represent the total number of experiments the authors have conducted to date.

Watch this Scientific Journal Video about Electroporation-mediated RNA Interference Method in Odonata at JoVE.com

Electroporation-mediated RNA Interference Method in Odonata

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, ...

electroporation-mediated-rna-interference-method-in-odonata

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

5.6K Views.  The University of Tokyo. 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).

Video: Electroporation-mediated RNA Interference Method in Odonata

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative without tools to study gene function. The development and sharing of detailed protocols for the establishment of new model systems in laboratories, and for tools to carry out functional studies, is thus crucial for leveraging the power of genomics. Coleoptera (beetles) are the largest clade of insects and occupy virtually all types of habitats on the planet. In addition to providing ideal models for fundamental research, studies of beetles can have impacts on pest control as they are often pests of households, agriculture, and food industries. Detailed protocols for rearing and maintenance of D. maculatus laboratory colonies and for carrying out dsRNA-mediated interference in D. maculatus are presented. Both embryonic and parental RNAi procedures-including apparatus set up, preparation, injection, and post-injection recovery-are described. Methods are also presented for analyzing embryonic phenotypes, including viability, patterning defects in hatched larvae, and cuticle preparations for unhatched larvae. These assays, together with in situ hybridization and immunostaining for molecular markers, make D. maculatus an accessible model system for basic and applied research. They further provide useful information for establishing procedures in other emerging insect model systems.

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Here, we present protocols for rearing an intermediate-germ beetle, Dermestes maculatus (D. maculatus) in the lab. We also share protocols for embryonic and parental RNAi and methods for analyzing embryonic phenotypes to study gene function in this species.

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative ...

Genetics

Double-stranded RNA Oral Delivery Methods to Induce RNA Interference in Phloem and Plant-sap-feeding Hemipteran Insects

Phloem and plant sap feeding insects invade the integrity of crops and fruits to retrieve nutrients, in the process damaging food crops. Hemipteran insects account for a number of economically substantial pests of plants that cause damage to crops by feeding on phloem sap. The brown marmorated stink bug (BMSB), Halyomorpha halys (Heteroptera: Pentatomidae) and the Asian citrus psyllid (ACP), Diaphorina citri Kuwayama (Hemiptera: Liviidae) are hemipteran insect pests introduced in North America, where they are an invasive agricultural pest of high-value specialty, row, and staple crops and citrus fruits, as well as a nuisance pest when they aggregate indoors. Insecticide resistance in many species has led to the development of alternate methods of pest management strategies. Double-stranded RNA (dsRNA)-mediated RNA interference (RNAi) is a gene silencing mechanism for functional genomic studies that has potential applications as a tool for the management of insect pests. Exogenously synthesized dsRNA or small interfering RNA (siRNA) can trigger highly efficient gene silencing through the degradation of endogenous RNA, which is homologous to that presented. Effective and environmental use of RNAi as molecular biopesticides for biocontrol of hemipteran insects requires the in vivo delivery of dsRNAs through feeding. Here we demonstrate methods for delivery of dsRNA to insects: loading of dsRNA into green beans by immersion, and absorbing of gene-specific dsRNA with oral delivery through ingestion. We have also outlined non-transgenic plant delivery approaches using foliar sprays, root drench, trunk injections as well as clay granules, all of which may be essential for sustained release of dsRNA. Efficient delivery by orally ingested dsRNA was confirmed as an effective dosage to induce a significant decrease in expression of targeted genes, such as juvenile hormone acid O-methyltransferase (JHAMT) and vitellogenin (Vg). These innovative methods represent strategies for delivery of dsRNA to use in crop protection and overcome environmental challenges for pest management.

This article demonstrates novel techniques developed for oral delivery of double-stranded RNA (dsRNA) through the vascular tissues of plants for RNA interference (RNAi) in phloem sap feeding insects.

Phloem and plant sap feeding insects invade the integrity of crops and fruits to retrieve nutrients, in the process damaging food crops. Hemipteran ...

Environment

Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

Synthetic dsRNAs, used to induce RNA interference, may have dose dependent phenotypic effects. These effects are difficult to define if the dsRNAs are delivered using a non-quantitative method. Accurate delivery of known quantities of nucleic acids or other chemicals is critical to measure the efficacy of the compound being tested and to allow reliable comparison between compounds.
Here we provide a reproducible, quantitative microinjection protocol that ensures accurate delivery of specific doses of dsRNA, reducing the mortality typically induced by injection injury. These modifications include the addition of Rhodamine B, a graduated injection needle, and an improved recovery method borrowed from Isoe and Collins. This method allows calculation of dose responses and facilitates comparisons between compounds. Versions of this method have been successfully used on three genera of mosquitoes as well as house flies to assess the reduction in fecundity resulting from gene silencing of ribosomal RNA transcripts.
This protocol provides strategies to reduce several challenges of small insect microinjection. Together, mechanical delivery of dsRNAs accompanied by visual verification, identification of effective locations for delivery, and inclusion of a post-injection recovery period ensure accurate dosing and low injury mortality. This protocol also describes an oviposition bioassay for uniform determination of effects on fecundity.

This protocol describes a microinjection methodology that we have standardized and used for several years to deliver specific quantities of nucleic acids directly to the hemolymph of mosquitoes and house flies. This protocol results in minimal injection mortality and allows dose correlated measurements of fecundity.

Synthetic dsRNAs, used to induce RNA interference, may have dose dependent phenotypic effects. These effects are difficult to define if the dsRNAs are ...

RNA Interference in Aquatic Beetles as a Powerful Tool for Manipulating Gene Expression at Specific Developmental Time Points

RNA interference (RNAi) remains a powerful technique that allows for the targeted reduction of gene expression through mRNA degradation. This technique is applicable to a wide variety of organisms and is highly efficient in the species-rich order Coleoptera (beetles). Here, we summarize the necessary steps for developing this technique in a novel organism and illustrate its application to the different developmental stages of the aquatic diving beetle Thermonectus marmoratus. Target gene sequences can be obtained cost-effectively through the assembly of transcriptomes against a close relative with known genomics or de novo. Candidate gene cloning utilizes a specific cloning vector (the pCR4-TOPO plasmid), which allows the synthesis of double-stranded RNA (dsRNA) for any gene with the use of a single common primer. The synthesized dsRNA can be injected into either embryos for early developmental processes or larvae for later developmental processes. We then illustrate how RNAi can be injected into aquatic larvae using immobilization in agarose. To demonstrate the technique, we provide several examples of RNAi experiments, generating specific knockdowns with predicted phenotypes. Specifically, RNAi for the tanning gene laccase2 leads to cuticle lightening in both larvae and adults, and RNAi for the eye pigmentation gene white produces a lightening/lack of pigmentation in eye tubes. In addition, the knockdown of a key lens protein leads to larvae with optical deficiencies and a reduced ability to hunt prey. Combined, these results exemplify the power of RNAi as a tool for investigating both morphological patterning and behavioral traits in organisms with only transcriptomic databases.

RNA interference is a widely applicable, powerful technique for manipulating gene expression at specific developmental stages. Here, we describe the necessary steps for implementing this technique in the aquatic diving beetle Thermonectus marmoratus, from the acquisition of gene sequences to the knockdown of genes that affect structure or behavior.

RNA interference (RNAi) remains a powerful technique that allows for the targeted reduction of gene expression through mRNA degradation. This technique is ...

Applications of RNA Interference in American Cockroach

Cockroaches, a sanitary pest, are essential species in insect developmental and metamorphic studies due to their easy feeding and hemimetabolous characteristics. Altogether with well-annotated genome sequences, these advantages have made American cockroach, Periplaneta americana, an important hemimetabolous insect model. Limited by the shortage of knockout strategy, effective RNA interference (RNAi)-based gene knockdown becomes an indispensable technique in functional gene research of P. americana. The present protocol describes the RNAi operation techniques in P. americana. The protocol includes (1) selection of the P. americana at proper developmental stages, (2) preparation for the injection setting, (3) dsRNA injection, and (4) gene knockdown efficiency detection. RNAi is a powerful reverse genetic tool in P. americana. The majority of P. americana tissues are sensitive to extracellular dsRNA. Its simplicity allows researchers to quickly obtain dysfunctional phenotypes under one or multiple targeting dsRNA injections, enabling researchers to better use the P. americana for developmental and metamorphic studies.

The present protocol describes step-by-step guidelines for the RNAi operation techniques in P. americana.

Cockroaches, a sanitary pest, are essential species in insect developmental and metamorphic studies due to their easy feeding and hemimetabolous ...

RNAi Interference by dsRNA Injection into Drosophila Embryos

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements and tools for genetic manipulation have become available in Drosophila 2. Soon after the initial discovery by Frie and Mello 3 that double stranded RNA can be used to knockdown the activity of individual genes in Caenorhabditis elegans, RNA interference (RNAi) was shown to provide a powerful reverse genetic approach to analyze gene functions in Drosophila organ development 4, 5.
Many organs, including lung, kidney, liver, and vascular system, are composed of branched tubular networks that transport vital fluids or gases 6, 7. The analysis of Drosophila tracheal formation provides an excellent model system to study the morphogenesis of other tubular organs 8. The Berkeley Drosophila genome project has revealed hundreds of genes that are expressed in the tracheal system. To study the molecular and cellular mechanism of tube formation, the challenge is to understand the roles of these genes in tracheal development. Here, we described a detailed method of dsRNA injection into Drosophila embryo to knockdown individual gene expression. We successfully knocked down endogenous dysfusion(dys) gene expression by dsRNA injection. Dys is a bHLH-PAS protein expressed in tracheal fusion cells, and it is required for tracheal branch fusion 9, 10. dys-RNAi completely eliminated dys expression and resulted in tracheal fusion defect. This relatively simple method provides a tool to identify genes requried for tissure and organ development in Drosophila.

RNAi Interference by dsRNA Injection into Drosophila Embryos

RNA interference has been proven very effective to analyze gene function in Drosophila tracheal development. A detailed protocol used by Jiang lab to inject dsRNA into fly embryos to knockdown gene expression is illustrated. This technique has the potential for screening genes required for tissue and organ development in Drosophila.

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements ...

Biology

Chitosan/Interfering RNA Nanoparticle Mediated Gene Silencing in Disease Vector Mosquito Larvae

Vector mosquitoes inflict more human suffering than any other organism&#8212;and kill more than one million people each year. The mosquito genome projects facilitated research in new facets of mosquito biology, including functional genetic studies in the primary African malaria vector Anopheles gambiae and the dengue and yellow fever vector Aedes aegypti. RNA interference- (RNAi-) mediated gene silencing has been used to target genes of interest in both of these disease vector mosquito species. Here, we describe a procedure for preparation of chitosan/interfering RNA nanoparticles that are combined with food and ingested by larvae. This technically straightforward, high-throughput, and relatively inexpensive methodology, which is compatible with long double stranded RNA (dsRNA) or small interfering RNA (siRNA) molecules, has been used for the successful knockdown of a number of different genes in A. gambiae and A. aegypti larvae. Following larval feedings, knockdown, which is verified through qRT-PCR or in situ hybridization, can persist at least through the late pupal stage. This methodology may be applicable to a wide variety of mosquito and other insect species, including agricultural pests, as well as other non-model organisms. In addition to its utility in the research laboratory, in the future, chitosan, an inexpensive, non-toxic and biodegradable polymer, could potentially be utilized in the field.

Here we describe a procedure for inhibiting gene function in disease vector mosquitoes through the use of chitosan/interfering RNA nanoparticles that are ingested by larvae.

Vector mosquitoes inflict more human suffering than any other organism&#8212;and kill more than one million people each year. The mosquito genome projects ...

RNAi Trigger Delivery into Anopheles gambiae Pupae

RNA interference (RNAi), a naturally occurring phenomenon in eukaryotic organisms, is an extremely valuable tool that can be utilized in the laboratory for functional genomic studies. The ability to knockdown individual genes selectively via this reverse genetic technique has allowed many researchers to rapidly uncover the biological roles of numerous genes within many organisms, by evaluation of loss-of-function phenotypes. In the major human malaria vector Anopheles gambiae, the predominant method used to reduce the function of targeted genes involves injection of double-stranded (dsRNA) into the hemocoel of the adult mosquito. While this method has been successful, gene knockdown in adults excludes the functional assessment of genes that are expressed and potentially play roles during pre-adult stages, as well as genes that are expressed in limited numbers of cells in adult mosquitoes. We describe a method for the injection of Serine Protease Inhibitor 2 (SRPN2) dsRNA during the early pupal stage and validate SRPN2 protein knockdown by observing decreased target protein levels and the formation of melanotic pseudo-tumors in SRPN2 knockdown adult mosquitoes. This evident phenotype has been described previously for adult stage knockdown of SRPN2 function, and we have recapitulated this adult phenotype by SRPN2 knockdown initiated during pupal development. When used in conjunction with a dye-labeled dsRNA solution, this technique enables easy visualization by simple light microscopy of injection quality and distribution of dsRNA in the hemocoel.

RNAi Trigger Delivery into Anopheles gambiae Pupae

RNA interference (RNAi) is an extremely valuable tool for uncovering gene function. However, the ability to target genes using RNAi during pre-adult stages is limited in the major human malaria vector Anopheles gambiae. We describe an RNAi protocol to reduce gene function via direct injection during pupal development.

RNA interference (RNAi), a naturally occurring phenomenon in eukaryotic organisms, is an extremely valuable tool that can be utilized in the laboratory ...

Practical Use of RNA Interference: Oral Delivery of Double-stranded RNA in Liposome Carriers for Cockroaches

RNA interference (RNAi) has been widely applied for uncovering the biological functions of numerous genes, and has been envisaged as a pest control tool operating by disruption of essential gene expression. Although different methods, such as injection, feeding, and soaking, have been reported for successful delivery of double-stranded RNA (dsRNA), the efficiency of RNAi through oral delivery of dsRNA is highly variable among different insect groups. The German cockroach, Blattella germanica, is highly sensitive to the injection of dsRNA, as shown by many studies published previously. The present study describes a method to demonstrate that the dsRNA encapsulated with liposome carriers is sufficient to retard the degradation of dsRNA by midgut juice. Notably, the continuous feeding of dsRNA encapsulated by liposomes significantly reduces the tubulin expression in the midgut, and led to the death of cockroaches. In conclusion, the formulation and utilization of dsRNA lipoplexes, which protect dsRNA against nucleases, could be a practical use of RNAi for insect pest control in the future.

This manuscript demonstrates the depletion of gene expression in the midgut of the German cockroach through oral ingestion of double-stranded RNA encapsulated in liposomes.

RNA interference (RNAi) has been widely applied for uncovering the biological functions of numerous genes, and has been envisaged as a pest control tool ...

Effective Oral RNA Interference (RNAi) Administration to Adult Anopheles gambiae Mosquitoes

RNA interference has been a heavily utilized tool for reverse genetic analysis for two decades. In adult mosquitoes, double-stranded RNA (dsRNA) administration has been accomplished primarily via injection, which requires significant time and is not suitable for field applications. To overcome these limitations, here we present a more efficient method for robust activation of RNAi by oral delivery of dsRNA to adult Anopheles gambiae. Long dsRNAs were produced in Escherichia coli strain HT115 (DE3), and a concentrated suspension of heat-killed dsRNA-containing bacteria in 10% sucrose was offered on cotton balls ad-libitum to adult mosquitoes. Cotton balls were replaced every 2 days for the duration of the treatment. Use of this method to target doublesex (a gene involved in sex differentiation) or fork head (which encodes a salivary gland transcription factor) resulted in reduced target gene expression and/or protein immunofluorescence signal, as measured by quantitative Real-Time PCR (qRT-PCR) or fluorescence confocal microscopy, respectively. Defects in salivary gland morphology were also observed. This highly flexible, user-friendly, low-cost, time-efficient method of dsRNA delivery could be broadly applicable to target genes important for insect vector physiology and beyond.

Effective Oral RNA Interference RNAi Administration to Adult Anopheles gambiae Mosquitoes

The oral administration of dsRNA produced by bacteria, a delivery method for RNA interference (RNAi) that is routinely used in Caenorhabditis elegans, was successfully applied here to adult mosquitoes. Our method allows for robust reverse genetics studies and transmission-blocking vector studies without the use of injection.

Electroporation-mediated RNA Interference Method in Odonata

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Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Double-stranded RNA Oral Delivery Methods to Induce RNA Interference in Phloem and Plant-sap-feeding Hemipteran Insects

Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

RNA Interference in Aquatic Beetles as a Powerful Tool for Manipulating Gene Expression at Specific Developmental Time Points

Applications of RNA Interference in American Cockroach

RNAi Interference by dsRNA Injection into Drosophila Embryos

Chitosan/Interfering RNA Nanoparticle Mediated Gene Silencing in Disease Vector Mosquito Larvae

RNAi Trigger Delivery into Anopheles gambiae Pupae

Practical Use of RNA Interference: Oral Delivery of Double-stranded RNA in Liposome Carriers for Cockroaches

Effective Oral RNA Interference (RNAi) Administration to Adult Anopheles gambiae Mosquitoes