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>

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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#181;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&#966;)</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&#160;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&#160;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&#160;Pure Chemical Corporation</td>
			<td>163-03545</td>
			<td>For PBS</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>FUJIFILM Wako&#160;Pure Chemical Corporation</td>
			<td>169-04245</td>
			<td>For PBS</td>
		</tr>
		<tr>
			<td>KOAc</td>
			<td>FUJIFILM Wako&#160;Pure Chemical Corporation</td>
			<td>160-03175</td>
			<td>For injection buffer</td>
		</tr>
		<tr>
			<td>KOH</td>
			<td>FUJIFILM Wako&#160;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)2</td>
			<td>FUJIFILM Wako&#160;Pure Chemical Corporation</td>
			<td>130-00095</td>
			<td>For injection buffer</td>
		</tr>
		<tr>
			<td>Na2HPO4</td>
			<td>FUJIFILM Wako&#160;Pure Chemical Corporation</td>
			<td>197-02865</td>
			<td>For PBS</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>FUJIFILM Wako&#160;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&#160;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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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

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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

Transcriptional Analysis by Nascent RNA FISH of In Vivo Trophoblast Giant Cells or In Vitro Short-term Cultures of Ectoplacental Cone Explants

The placenta derives from one extra-embryonic lineage, the trophectoderm. In the peri-implantation murine blastocyst, mural trophectoderm cells differentiate into primary trophoblast giant cells (TGCs) while the polar trophectoderm overlying the inner cell mass continues to proliferate later differentiating into secondary TGCs. TGCs play a key role in developing placenta and are essential for a successful pregnancy. Investigation of transcriptional regulation of specific genes during post-implantation development can give insights into TGCs development. Cells of the ectoplacental cone (EPC) from embryos at 7-7.5 days of gestation (E7-7.5), derived from the polar trophectoderm, differentiate into secondary TGCs1. TGCs can be studied in situ, on cryostat sections of embryos at E7 although the number of TGCs is very low at this stage. An alternative means of analyzing secondary TGCs is to use short-term cultures of individual EPCs from E7 embryos. We propose a technique to investigate the transcriptional status of genes of interest both in vivo and in vitro at the single-cell level using fluorescent in situ hybridization (RNA FISH) to visualize nascent transcripts. This technique provides a direct readout of gene expression and enables assessment of the chromosomal status of TGCs, which are large endoreplicating cells. Indeed, a key feature of terminal differentiation of TGCs is that they exit the cell cycle and undergo multiple rounds of endoreplication.This approach can be applied to detect expression of any gene expressed from autosomes and/or sex chromosomes and can provide important information into developmental mechanisms as well as placental diseases.

Transcriptional  Analysis by Nascent RNA FISH of In Vivo Trophoblast Giant Cells or In Vitro Short-term Cultures of Ectoplacental Cone Explants

Trophoblast giant cells (TGCs) play a key role in the placenta to ensure a healthy pregnancy. We present a protocol for assessing the transcriptional status of genes in TGCs by nascent fluorescent in situ hybridization on cryostat sections of post-implantation embryos or short-term cultures of embryonic day 7 ectoplacental cones.

The placenta derives from one extra-embryonic lineage, the trophectoderm. In the peri-implantation murine blastocyst, mural trophectoderm cells ...

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

The evolutionarily conserved&#160;extracellular signal transducing RTK-RAS-ERK pathway is an important kinase-signaling cascade that controls multiple cellular and developmental processes principally via activation of ERK, the terminal kinase of the pathway. Tight regulation of ERK activity is essential for normal development and homeostasis; overly active ERK results in excessive cellular proliferation, while underactive ERK causes cell death. C. elegans is a powerful model system that has helped characterize the function and regulation of RTK-RAS-ERK signaling pathway during development. In particular, the RTK-RAS-ERK pathway is essential for C. elegans germline development, which is the focus of this method. Using antibodies specific to the active, diphosphorylated form of ERK (dpERK), the stereotypical localization pattern can be visualized within the germline. Because this pattern is both spatially and temporally controlled, the ability to reproducibly assay dpERK is useful to identify regulators of the pathway that affect dpERK signal duration and amplitude and thus germline development. Here we demonstrate how to successfully dissect, stain, and image dpERK within the C. elegans gonad. This method can be adapted for spatial localization of any signaling or structural protein in the C. elegans gonad, provided an antibody compatible with immunofluorescence is available.

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

We present an immunofluorescence&#160;imaging-based method for spatial and temporal localization of active ERK in the dissected C. elegans gonad. The protocol described here can be adapted for visualization of any signaling or structural protein in the C. elegans gonad, provided a suitable antibody reagent is available.

The evolutionarily conserved&#160;extracellular signal transducing RTK-RAS-ERK pathway is an important kinase-signaling cascade that controls multiple ...

Directed Differentiation of Primitive and Definitive Hematopoietic Progenitors from Human Pluripotent Stem Cells

One of the major goals for regenerative medicine is the generation and maintenance of hematopoietic stem cells (HSCs) derived from human pluripotent stem cells (hPSCs). Until recently, efforts to differentiate hPSCs into HSCs have predominantly generated hematopoietic progenitors that lack HSC potential, and instead resemble yolk sac hematopoiesis.&#160;These resulting hematopoietic progenitors may have limited utility for in vitro disease modeling of various adult hematopoietic disorders, particularly those of the lymphoid lineages. However, we have recently described methods to generate erythro-myelo-lymphoid multilineage definitive hematopoietic progenitors from hPSCs using a stage-specific directed differentiation protocol, which we outline here. Through enzymatic dissociation of hPSCs on basement membrane matrix-coated plasticware, embryoid bodies (EBs) are formed. EBs are differentiated to mesoderm by recombinant BMP4, which is subsequently specified to the definitive hematopoietic program by the GSK3&#946; inhibitor, CHIR99021. Alternatively, primitive hematopoiesis is specified by the PORCN inhibitor, IWP2. Hematopoiesis is further driven through the addition of recombinant VEGF and supportive hematopoietic cytokines. The resulting hematopoietic progenitors generated using this method have the potential to be used for disease and developmental modeling, in vitro.

Here, we present human pluripotent stem cell (hPSC) culture protocols, used to differentiate hPSCs into CD34+ hematopoietic progenitors. This method uses stage-specific manipulation of canonical WNT signaling to specify cells exclusively to either the definitive or primitive hematopoietic program.

One of the major goals for regenerative medicine is the generation and maintenance of hematopoietic stem cells (HSCs) derived from human pluripotent stem ...

A Method for Characterizing Embryogenesis in Arabidopsis

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, early stage plant embryos are small and contain few cells, making them difficult to observe and analyze. A method is described here for characterizing pattern formation in plant embryos under a microscope using the model organism Arabidopsis. Following the clearance of fresh ovules using Hoyer's solution, the cell number in and morphology of embryos could be observed, and their developmental stage could be determined by differential interference contrast microscopy using a 100X oil immersion lens. In addition, the expression of specific marker proteins tagged with Green Fluorescent Protein (GFP) was monitored to annotate cell identity specification during embryo patterning by confocal laser scanning microscopy. Thus, this method can be used to observe pattern formation in wild-type plant embryos at the cellular and molecular levels, and to characterize the role of specific genes in embryo patterning by comparing pattern formation in embryos from wild-type plants and embryo-lethal mutants. Therefore, the method can be used to characterize embryogenesis in Arabidopsis.

A Method for Characterizing Embryogenesis in Arabidopsis

This protocol outlines a method for observing embryogenesis in Arabidopsis via ovule clearance followed by the inspection of embryo pattern formation under a microscope.

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by Sertoli cell junctions at the blood-testis barrier (BTB), which is the tightest tissue barrier in the mammalian body and segregates the seminiferous epithelium into two compartments, a basal and an adluminal. The BTB creates a unique microenvironment for germ cells in meiosis I/II and for the development of postmeiotic spermatids into spermatozoa via spermiogenesis. Here, we describe a reliable assay to monitor BTB integrity of mouse testis in vivo. An intact BTB blocks the diffusion of FITC-conjugated inulin from the basal to the apical compartment of the seminiferous tubules. This technique is suitable for studying gene candidates, viruses, or environmental toxicants that may affect BTB function or integrity, with an easy procedure and a minimal requirement of surgical skills compared to alternative methods.

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Here, we present a protocol to assess the blood-testis barrier integrity by injecting inulin-FITC into testes. This is an efficient in vivo method to study blood-testis barrier integrity that can be compromised by genetic and environmental elements.

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by ...

RNA-based Reprogramming of Human Primary Fibroblasts into Induced Pluripotent Stem Cells

Induced pluripotent stem cells (iPSCs) have proven to be a valuable tool to study human development and disease. Further advancing iPSCs as a regenerative therapeutic requires a safe, robust, and expedient reprogramming protocol. Here, we present a clinically relevant, step-by-step protocol for the extremely high-efficiency reprogramming of human dermal fibroblasts into iPSCs using a non-integrating approach. The core of the protocol consists of expressing pluripotency factors (SOX2, KLF4, cMYC, LIN28A, NANOG, OCT4-MyoD fusion) from in vitro transcribed messenger RNAs synthesized with modified nucleotides (modified mRNAs). The reprogramming modified mRNAs are transfected into primary fibroblasts every 48 h together with mature embryonic stem cell-specific microRNA-367/302 mimics for two weeks. The resulting iPSC colonies can then be isolated and directly expanded in feeder-free conditions. To maximize efficiency and consistency of our reprogramming protocol across fibroblast samples, we have optimized various parameters including the RNA transfection regimen, timing of transfections, culture conditions, and seeding densities. Importantly, our method generates high-quality iPSCs from most fibroblast sources, including difficult-to-reprogram diseased, aged, and/or senescent samples.

Here we describe a clinically relevant, high-efficiency, feeder-free method to reprogram human primary fibroblasts into induced pluripotent stem cells using modified mRNAs encoding reprogramming factors and mature microRNA-367/302 mimics. Also included are methods to assess reprogramming efficiency, expand clonal iPSC colonies, and confirm expression of the pluripotency marker TRA-1-60.

Induced pluripotent stem cells (iPSCs) have proven to be a valuable tool to study human development and disease. Further advancing iPSCs as a regenerative ...

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

The Drosophila pupal retina provides an excellent model system for the study of morphogenetic processes during development. In this paper, we present a reliable protocol for the dissection of the delicate Drosophila pupal retina. Our surgical approach utilizes readily-available microdissection tools to open pupae and precisely extract eye-brain complexes. These can be fixed, subjected to immunohistochemistry, and retinas then mounted onto microscope slides and imaged if the goal is to detect cellular or subcellular structures. Alternatively, unfixed retinas can be isolated from brain tissue, lysed in appropriate buffers and utilized for protein gel electrophoresis or mRNA extraction (to assess protein or gene expression, respectively). Significant practice and patience may be required to master the microdissection protocol described, but once mastered, the protocol enables relatively quick isolation of mainly undamaged retinas.

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

This paper presents a surgical method for dissecting Drosophila pupal retinas along with protocols for the processing of tissue for immunohistochemistry, western analysis, and RNA-extraction.&#160;

The Drosophila pupal retina provides an excellent model system for the study of morphogenetic processes during development. In this paper, we present a ...

Electroporation Method for In Vivo Delivery of Plasmid DNA in the Adult Zebrafish Telencephalon

Electroporation is a transfection method in which an electrical field is applied to cells to create temporary pores in a cell membrane and increase its permeability, thereby allowing different molecules to be introduced to the cell. In this paper, electroporation is used to introduce plasmids to ependymoglial cells, which line the ventricular zone of the adult zebrafish telencephalon. A fraction of these cells shows stem cell properties and generates new neurons in the zebrafish brain; therefore, studying their behavior is essential to determine their roles in neurogenesis and regeneration. The introduction of plasmids via electroporation enables long-term labeling and tracking of a single ependymoglial cell. Furthermore, plasmids such as Cre recombinase or Cas9 can be delivered to single ependymoglial cells, which enables gene recombination or gene editing and provides a unique opportunity to assess the cell&#8217;s autonomous gene function in a controlled, natural environment. Finally, this detailed, step-by-step electroporation protocol is used to obtain successful introduction of plasmids into a large number of single ependymoglial cells.

Presented here is an electroporation method for plasmid DNA delivery and ependymoglial cell labeling in the adult zebrafish telencephalon. This protocol is a quick and efficient method to visualize and trace individual ependymoglial cells and opens up new possibilities to apply electroporation to a broad range of genetic manipulations.

Electroporation is a transfection method in which an electrical field is applied to cells to create temporary pores in a cell membrane and increase its ...

Electroporation-mediated RNA Interference Method in Odonata

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Chapters in this video

Transcriptional Analysis by Nascent RNA FISH of In Vivo Trophoblast Giant Cells or In Vitro Short-term Cultures of Ectoplacental Cone Explants

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

Directed Differentiation of Primitive and Definitive Hematopoietic Progenitors from Human Pluripotent Stem Cells

A Method for Characterizing Embryogenesis in Arabidopsis

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

RNA-based Reprogramming of Human Primary Fibroblasts into Induced Pluripotent Stem Cells

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

Electroporation Method for In Vivo Delivery of Plasmid DNA in the Adult Zebrafish Telencephalon