We thank Drs. Alison Heffer and Yong Lu for setting up the microinjection apparatus and sharing their invaluable knowledge and experience with insect RNAi. This work was supported by the National Institutes of Health (R01GM113230 to L.P.).

1. Rearing of D. maculatus
NOTE: A breeding colony of D. maculatus was set up in the authors&#39; lab using adults and larvae purchased commercially. The species identity was verified using DNA barcoding53.
<ol>
	<li>To set up a new cage in the lab, spread a thin layer of wood shavings into a medium-size insect cage (30.5 x 19 x 20.3 cm3). Place a ~ 10 x 6 x 3 cm3 chunk of Styrofoam in the cage to let larvae hide for pupation. Add 20 - 50 beetles (either adults or late instar larvae). Beetles will hide in the wood shavings.</li>
	<li>Add wet cat food in a Petri dish or a weighing boat. Cover the cage with mesh cloth and place the cage in an incubator.</li>
	<li>For maintaining a breeding colony in the lab, set the temperature between 25 and 30 &#176;C. D. maculatus grows faster at higher temperatures, but this also promotes the growth of fungus and mites. To maintain a healthy colony, use 25 - 28 &#176;C for regular stock maintenance and 30 &#176;C for rapidly expanding the colony. The life cycle takes approximately three weeks to four months depending on environmental factors50 (Figure 2).</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/54976/54976fig2.jpg" /> 
Figure 2: D. maculatus Lab Colony. Photograph of a typical D. maculatus insect cage is shown. Wood shavings are spread to let the beetles hide. Cat food is added in a small Petri dish or weighing boat. Styrofoam is placed into the cage as a refuge for final instar larvae to pupate. The cage shown here is 30.5 x 19 x 20.3 cm3 and houses a few hundred larvae, pupae, and adults. <a href="http://cloudfront.jove.com/files/ftp_upload/54976/54976fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="4">
	<li>Leave eggs in the cage to mature. To expand the colony, establish new cages with eggs (collected as described in Section 2), larvae, or adults, as above. D. maculatus lay eggs throughout the cage, especially near the food source.</li>
	<li>Replace wet cat food about twice a week. 
	NOTE: Due to the unpleasant odor of wet cat food, alternative food sources were also tested (see Discussion).</li>
</ol>
2. Embryo Collection
<ol>
	<li>To set up a collection, separate at least 50 males and 50 females from the colony. Young adults are best. Place adults in a mini-sized cage (17.8 x 10.2 x 12.7 cm3) without wood shavings. Add cat food in a weighing boat.</li>
	<li>Before starting collection, check the cage carefully for any embryos present and remove them. Put a stretched cotton ball into the cage, and leave the cage in a 30 &#176;C incubator. After the appropriate time window, the cotton ball will be ready for embryo collection.</li>
	<li>Fold a piece of black construction paper (A4 size) in half to create a crease, and then unfold.</li>
	<li>Remove the stretched cotton ball from the cage. Remove adults from the cotton ball and put them back into the cage.</li>
	<li>While pinching the stretched cotton ball very gently, tear it apart slowly into thin cotton filaments to let the eggs fall onto the black paper. 
	NOTE: D. maculatus embryos are fragile and hard to see in cotton. They can be easily crushed if holding the cotton ball too firmly.</li>
</ol>
3. dsRNA Preparation
<ol>
	<li>DNA Template Preparation for dsRNA Synthesis
	<ol>
		<li>Run 4 - 6 50 &#181;l PCR reactions using primers containing T7 promoter sequences at both 5&#39; ends to amplify DNA template according to manufacturer&#39;s protocol.</li>
		<li>Purify PCR product using a commercial PCR purification kit, following manufacturer&#39;s instructions. The ideal final concentration of DNA template is &#8805; 100 ng/&#181;l.</li>
	</ol>
	</li>
	<li>Injection Buffer Preparation
	<ol>
		<li>Prepare 100 ml of injection buffer (0.1 mM NaH2PO4, 5 mM KCl). Adjust pH to 6.8 with 5 M NaOH.</li>
		<li>Store aliquots at -20 &#176;C.</li>
	</ol>
	</li>
	<li>dsRNA Synthesis
	<ol>
		<li>Set up a reaction in a 0.2 ml PCR tube on ice and carry out an in vitro transcription reaction with T7 polymerase, following manufacturer&#39;s instructions, using ~ 500 ng template per reaction.</li>
		<li>Incubate the tube at 37 &#176;C overnight.</li>
		<li>Digest the DNA template with DNase, following manufacturer&#39;s instructions.</li>
		<li>Heat the tube to 94 &#176;C and hold at 94 &#176;C for 3 min.</li>
		<li>To anneal the dsRNA, slowly cool the tube by 1 &#176;C/min until it reaches 45 &#176;C after 1 hr. 
		NOTE: Steps 3.3.2 through 3.3.5 can be performed in a thermocycler.</li>
		<li>To ethanol precipitate&#160;of dsRNA, add 280 &#181;l of RNase free water to 20 &#181;l of reaction to adjust to a final volume of 300 &#181;l. Add 30 &#181;l of 3 M NaOAc (pH 5.2) and 650 &#181;l of ethanol. Place tube in -20 &#176;C freezer overnight.</li>
		<li>After centrifuging at 15,682 x g for 30 min at 4 &#176;C, wash the dsRNA pellet with 70% ethanol. Dry pellet and dissolve in 20 &#181;l injection buffer.</li>
		<li>Measure the concentration of the dsRNA using a spectrophotometer at A260. The ideal concentration is 3 - 5 &#181;g/&#181;l. Check the quality by running 5 &#181;l of a 1:25 dilution of dsRNA on a 1% agarose gel at 90 V for 30 min. A single band of expected size will be readily visible.</li>
		<li>Store the dsRNA at -20 &#176;C or colder. 
		NOTE: For every RNAi knockdown experiment, include a control dsRNA, which would not be expected to impact the process being studied. gfp dsRNA is an excellent control for most experiments. Prepare and inject the control dsRNA side-by-side with the experimental dsRNA.</li>
	</ol>
	</li>
</ol>
4. Assembly of Microinjector and Micromanipulator
NOTE: See Figure 3.
<ol>
	<li>Connect nitrogen or other air supply to the pressure input of a pneumatic pump instrument. If a pneumatic pump is not available, apply pressure using a 50 ml syringe connected to fine tubing.</li>
	<li>Connect a foot switch to the remote connector of the pump to control the eject pressure flow.</li>
	<li>Attach a glass capillary holder to the eject pressure port of the pump with tube.</li>
	<li>Fix the capillary holder on a micromanipulator close to a dissecting microscope.</li>
</ol>
5. Embryonic RNAi
NOTE: Figure 4 shows a flowchart of D. maculatus embryonic RNAi.
<ol>
	<li>eRNAi Preparation
	<ol>
		<li>Prepare an embryo collecting Petri dish lid (90 mm diameter). Place a piece of black filter paper to cover the inside of the lid. Put a standard microscope slide on the black paper.</li>
		<li>Dilute dsRNA to appropriate concentration with injection buffer. Use 2 - 3 &#181;g/&#181;l for initial experiments. Always keep dsRNA on ice prior to injection.</li>
		<li>Add food coloring at 1:40 dilution to the dsRNA and pipette gently several times to mix well.</li>
	</ol>
	</li>
	<li>Collecting Embryos for Injection 
	NOTE: At 25 &#176;C, nuclei in embryos 6 - 8 hr After Egg Laying (AEL) migrate towards the egg periphery53. A cellular blastoderm is established within 8 - 10 hr AEL53. To ensure embryos prior to cellular blastoderm stage are used for injection, 0 - 3 hr AEL embryos are recommended for use53. Embryo collection, preparation, and injection usually take less than 2 hr if the glass capillaries are in good shape. Therefore, the whole process can be completed within 5 hr AEL.
	<ol>
		<li>Collect embryos as described in steps 2.2-2.5.</li>
		<li>Tap the black construction paper to transfer embryos into collecting Petri dish lid.</li>
		<li>Stick double-sided tape along the edge of the slide.</li>
		<li>Using a paint brush, align the embryos on the tape perpendicular to the slide edge with the anterior or posterior end at the edge. Note that the anterior and posterior ends of early embryos are not readily distinguishable, thus on average half will be injected at the posterior end which is ideal.</li>
	</ol>
	</li>
	<li>Loading the dsRNA into the Glass Capillary
	<ol>
		<li>Take up 2 - 4 &#181;l of dsRNA into a 20 &#181;l microloader pipette tip.</li>
		<li>Insert the tip into a pre-pulled glass capillary (referred to as needle) and gently pipette the dsRNA solution into the capillary. Prepare needles from commercial glass capillaries using a micropipette puller. An example of an ideal pulled needle is shown in Figure 5. The length and taper of needles may need to be optimized after injection trials.</li>
		<li>Fix the glass capillary into the glass capillary holder.</li>
		<li>Assemble the capillary holder into the micromanipulator.</li>
	</ol>
	</li>
	<li>Injecting dsRNA into Embryos
	<ol>
		<li>Carefully transfer the slide with embryos onto the stage of the dissecting microscope.</li>
		<li>Move the slide to bring one embryo to the center of the field and focus microscope.</li>
		<li>Position the tip of the capillary with the micromanipulator while looking into the dissecting microscope. Bring the tip close to the end of the first embryo in the row.</li>
		<li>Switch on the nitrogen or other air supply.</li>
		<li>Set the solenoid input selector switch to vacuum.</li>
		<li>Adjust the eject pressure regulator to 10 - 15 psi. Adjust the pressure depending on the apparatus.</li>
		<li>Open the capillary if necessary. Once the tip is open, the colored dsRNA solution will fill the tip.</li>
		<li>Move the capillary tip forward to puncture the embryo. If the pressure setting is ideal, no embryonic fluid will flow into the capillary.</li>
		<li>Step on the foot switch to eject dsRNA solution into the embryo until an appropriate amount of solution is ejected. Figure 5 shows examples of embryo morphology after appropriate and inappropriate amounts of solution have been injected.</li>
		<li>Keep the capillary tip inside the embryo for ~ 2 sec, and then remove it.</li>
		<li>Move the capillary to the next embryo and repeat steps 5.4.8 - 5.4.10 until all embryos are injected. Change glass capillary if it gets clogged or breaks.</li>
	</ol>
	</li>
	<li>Post-injection Recovery and Incubation
	<ol>
		<li>After injection, place the slide in a Petri dish.</li>
		<li>Add a wet cotton ball to the Petri dish. Do not let the cotton ball touch the slide.</li>
		<li>Cover the Petri dish and wrap it with sealing film. Label the dish with the name of the dsRNA, concentration, date, time, and number of injected embryos.</li>
		<li>Place the Petri dish in a 30 &#176;C incubator until the embryos hatch.</li>
	</ol>
	</li>
</ol>
6. Parental RNAi
<ol>
	<li>Isolating Pupae for pRNAi
	<ol>
		<li>Collect pupae from the colony.</li>
		<li>Sort male and female pupae under microscope (see Figure 6). Keep them in two separate Petri dishes in a 30 &#176;C incubator to ensure that no mating occurs prior to injection.</li>
		<li>Check every day for eclosed adults. Pupation usually takes 5 - 7 days at 30 &#176;C.</li>
		<li>Transfer eclosed males and females (see Figure 6) to two separate Petri dishes and feed them cat food every other day until they are ready for injection.</li>
		<li>Proceed to injection once there are enough females and males at appropriate age. Females 4 to 8 days post-eclosion are best. For a typical analysis, 8 - 12 females are used. An equal number of males are needed for mating54.</li>
	</ol>
	</li>
	<li>Injecting dsRNA into Female Abdomen
	<ol>
		<li>Prepare dsRNA on ice (5.1.2 and 5.1.3).</li>
		<li>Attach a 32-gauge needle to a 10 &#181;l syringe.</li>
		<li>Anaesthetize females on CO2 stage.</li>
		<li>Load dsRNA solution into the syringe without taking up any air.</li>
		<li>Hold a female ventral side up with one hand and hold the syringe with the other.</li>
		<li>Gently penetrate the segmacoria (membrane) between sternites 2 and 3 with the tip of the needle. Figure 7 shows a female during injection. If the needle does not penetrate the tissue easily, angle the needle upward and slowly press down. Insert approximately 2 mm of the needle into the body.</li>
		<li>Check the syringe scale while slowly pushing the plunger, injecting ~ 2 &#181;l per female.</li>
		<li>After injecting, hold the needle still for at least 5 sec before removing it from the female&#39;s abdomen.</li>
		<li>Transfer the injected females to a Petri dish (90 mm diameter) and feed with cat food.</li>
		<li>Label the Petri dish with the dsRNA name, amount injected, date, and number of females.</li>
	</ol>
	</li>
	<li>Post-injection Recovery and Mating
	<ol>
		<li>Incubate the Petri dish in a 30 &#176;C incubator.</li>
		<li>After 24 hr, transfer injected females to a mini cage.</li>
		<li>Add to the cage an equal number of uninjected young males, and label the cage.</li>
		<li>Add a weighing boat with cat food into the cage.</li>
		<li>Leave the cage in a 30 &#176;C incubator to allow mating. After 24 hr, females should start to lay eggs and embryos can be collected daily or at required time intervals.</li>
	</ol>
	</li>
</ol>
7. Phenotypic Analysis after RNAi
NOTE: At 30 &#176;C, it takes ~ 55 hr for eggs to hatch50.
<ol>
	<li>eRNAi Viability Analysis
	<ol>
		<li>Calculate hatch rate by comparing the number of hatched larvae to the total number of injected embryos. Be sure to include a negative control injection as embryos may be harmed or killed by the injection procedure. Typical hatch rates vary from 30% - 60%. Beware of hatched larvae eating unhatched eggs.</li>
	</ol>
	</li>
	<li>pRNAi Viability Analysis 
	NOTE: If female viability is impacted by the gene targeted by RNAi, females injected with specific dsRNA but not negative control dsRNA will begin to die. If egg production or egg laying is impacted, females will exhibit partial or complete sterility. Score female survival compared to negative controls or compare total number of eggs laid by experimental and control females (7.2.3).
	<ol>
		<li>Set up embryo collection following step 2.2.</li>
		<li>After 24 hr, collect embryos following steps 2.4 - 2.5.</li>
		<li>Count the total number of the embryos.</li>
		<li>Transfer the embryos to a 90 mm Petri dish. Label the Petri dish with the gene targeted, collection date, and number of embryos.</li>
		<li>Incubate the embryos in a 30 &#176;C incubator until they hatch. Since hatched larvae feed on unhatched embryos, check the Petri dish frequently and separate hatched larvae from unhatched embryos using forceps (see Discussion).</li>
		<li>Count the number of hatched embryos and calculate the hatch rate.</li>
	</ol>
	</li>
	<li>Examining Cuticle Defects in Hatched Larvae
	<ol>
		<li>Collect hatched larvae in a 1.5 ml microcentrifuge tube.</li>
		<li>In a fume hood, add 1 ml of Fixation Solution.</li>
		<li>Leave the microcentrifuge tube at 4 &#176;C at least overnight to let the solution penetrate the larval tissue.</li>
		<li>To visualize, remove as much fixative from the tube as possible.</li>
		<li>Rinse larvae at least three times with PBST.</li>
		<li>Transfer larvae to a multi-well glass plate with PBST using a P-1000 pipette tip with the end cut off to widen it.</li>
		<li>Under a dissecting microscope, stretch larvae using forceps to examine defects. It is critical to stretch out larvae to examine defects as their bodies contract in fixative.</li>
	</ol>
	</li>
	<li>Examining Cuticle Defects in Unhatched Larvae 
	Note: This procedure is useful for examination of early embryonic phenotypes, especially for embryos that do not survive to hatching.
	<ol>
		<li>For pRNAi, add PBST into the Petri dish (7.2.4) and transfer unhatched embryos to a 1.5 ml microcentrifuge tube using a P-1000 pipette tip with the end cut off. For eRNAi, add PBST into the Petri dish (5.5.4), brush unhatched embryos off the microscope slide and transfer them to a microcentrifuge tube.</li>
		<li>Fix the embryos with Fixation Solution and rinse them with PBST for visualization following steps 7.3.2 - 7.3.6.</li>
		<li>Using forceps, dissect embryos out of the eggshell under a dissecting microscope in PBST.</li>
		<li>Transfer the embryos back to 1.5 ml microcentrifuge tube (~ 200 &#181;l of embryos in each tube).</li>
		<li>Remove as much solution as possible.</li>
		<li>Add 1 ml of 90% lactic acid/10% EtOH. Leave the centrifuge tube in a 60 &#176;C incubator at least overnight. 
		NOTE: Embryos can be left at 60 &#176;C for several days.</li>
		<li>To mount the embryos, pipette them onto a microscope slide using a P-1000 pipette tip with the end cut off.</li>
		<li>Manually position the embryos with forceps to an ideal orientation.</li>
		<li>Cover the embryos with a cover-slip and visualize under microscope using DIC.</li>
	</ol>
	</li>
	<li>Examining Cellular and Molecular Defects 
	NOTE: This procedure is useful for investigating the underlying causes of cuticle phenotypes or lethality that is not accompanied by cuticle defects.
	<ol>
		<li>For pRNAi, separate embryos from cotton balls at appropriate development stage and transfer them into an embryo collecting basket. The collection basket can be the same or similar to that used for Drosophila embryo collections.
		<ol>
			<li>Make the egg basket from a 25 ml plastic scintillation vial with the bottom of the vial removed, and a circle cut out of the lid (roughly 1 cm diameter). Place a circle of super fine mesh about 2.5 cm in diameter inside the lid, and then screw the scintillation vial on and use upside-down.</li>
			<li>As the mesh can be easily removed once the embryos are washed, immerse the mesh in a microcentrifuge tube with water to recover any embryos sticking to the mesh.</li>
		</ol>
		</li>
		<li>For eRNAi, at appropriate stage, add PBST to the Petri dish. Brush the embryos off the microscope slide and transfer them to an embryo collecting basket. 
		NOTE: Fixation, in situ hybridization, antibody staining, and nuclear staining protocols can be found in Xiang et al. 201553.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

While a small number of sophisticated model systems (mice, flies, worms) were developed during the 20th century, the 21st century has seen a wave of new animal systems being developed in labs throughout the world. These new systems allow scientists to address comparative, evolutionary questions that cannot be probed using only the &#39;standard&#39; model systems. This deployment of new models requires the rapid development of methods for lab culturing, gene identification, and functional approaches...

In 1998,&#160;Fire and Mello reported that double-stranded RNA (dsRNA) can induce inhibition of gene function in Caenorhabditis elegans1. This response triggered by dsRNA was named RNA interference (RNAi), and such RNAi-mediated gene silencing was reported to be conserved in animals, plants, and fungi2-7. In plants and some animals, RNAi functions systemically, meaning that the effect can spread to other cells/tissues where dsRNA is not directly introduced (reviewed in8-10). Scientists have made use of this endogenous cellular RNAi response by designing dsRNAs to target genes of interest, thereby knocking down gene function without directly manipulating the genome (reviewed in11-14).
RNAi is a powerful tool for functional studies due to the following advantages. First, even with minimal gene sequence information, a gene can be targeted using RNAi. This is especially important for studies of non-model organisms lacking genomic or transcriptomic data. Second, in organisms where the RNAi response is robustly systemic, RNAi-mediated gene knockdown can be performed at almost any developmental stage. This feature is very useful for studying the function of pleiotropic genes. Third, in some cases, RNAi effects spread to the gonads and progeny, such that phenotypes are observed in offspring15,16. This phenomenon, known as parental RNAi (pRNAi), is especially advantageous for genes impacting embryonic development, as numerous offspring produced by a single injected parent can be examined without direct manipulation of eggs. For these reasons, pRNAi is the method of choice. However, if pRNAi is ineffective, for example for genes required for oogenesis, then embryonic RNAi (eRNAi) must be used. Fourth, RNAi can be used to generate the equivalent of an allelic series in that the amount of dsRNA delivered can be varied over a range to produce weak to strong defects. Such a gradation of phenotypes can be helpful for understanding gene function when the gene is involved in a complex process and/or complete loss of function is lethal. Fifth, delivery of dsRNA is generally easy and feasible, especially in animals showing robust systemic RNAi responses. dsRNA can be introduced by microinjection1,5, feeding/ingestion17,18, soaking,19,20 and virus/bacteria-mediated delivery21,22. Sixth, unlike some gene targeting/editing methods, there is no need to screen for organisms carrying the mutation or to carry out genetic crosses to generate homozygotes when using RNAi. Therefore, compared to many other techniques for studying gene function, RNAi is fast, inexpensive, and can be applied for large-scale screens23-25.
The broad utility of RNAi provides means&#160;to carry out functional studies in a wide range of organisms, expanding the range of species available for study beyond the traditional model systems for which genetic tools have been developed. For example, studies using non-model systems are required to give insights into the evolution of genes and gene networks by comparing the functions of orthologs from species representing different development modes or exhibiting distinct morphological features26-29. These types of studies will provide a better understanding of biological diversity, with impacts for both applied and basic research.
Being the largest animal group on the planet, insects provide a great opportunity to explore the mechanisms underlying diversity. Additionally, insects are generally small, have short life cycles, high fecundity, and are easy to rear in the lab. In the past two decades, RNAi has been successfully applied in insects spanning orders, including Diptera (true flies)5, Lepidoptera (butterflies and moths)30, Coleoptera (beetles)16,31, Hymenoptera (sawflies, wasps, ants and bees)32, Hemiptera (true bugs), Isoptera (termites)34, Blattodea (cockroaches)35, Orthoptera (crickets, grasshoppers, locusts, and katydids)36and Phthiraptera (lice)37. Successful application of RNAi has provided functional data for studies of patterning in early embryogenesis (anterior-posterior axis32, dorsal-ventral axis28, segmentation26,38), sex determination39,40, chitin/cuticle biosynthesis41, ecdysone signaling42, social behavior43, and more. RNAi methods developed for different insect species may be of additional benefit in that they are likely to be useful for pest control (reviewed in44-46). RNAi effects will be gene-specific as well as species-specific, as long as non-conserved regions are chosen for targeting. For beneficial insect species like honeybees and silkworms, targeting genes vital for the survival of viruses or parasites to control infection may provide a novel strategy to protect these species47,48.
Dermestes maculatus (D. maculatus), common name hide beetle, is distributed worldwide except for Antarctica. As a holometabolous insect, the D. maculatus life cycle includes embryonic, larval, pupal, and adult stages (Figure 1). Because it feeds on flesh, D. maculatus is used in museums to skeletonize dead animals and forensic entomologists can use it to estimate time of death49,50. D. maculatus feeds on animal products including carcasses, dried meat, cheese, and the pupae/cocoons of other insects and thus causes damage to households, stored food, and the silk, cheese, and meat industries 51,52. Applying RNAi in this beetle could provide an efficient and environmentally friendly way to minimize its economic impact. Our lab has used D. maculatus as a new model insect to study segmentation53. In addition to being amenable to lab rearing, D. maculatus is of interest for basic research as it is an intermediate-germ developer, making it a useful species to study the transition between short- and long-germ development.
<img alt="Figure 1" src="/files/ftp_upload/54976/54976fig1.jpg" /> 
Figure 1: Life Cycle of D. maculatus. Photographs of D. maculatus at different life stages, as indicated. The life cycle from egg to adult takes three weeks at 30 &#176;C but longer at lower temperatures. (A, F) Freshly laid embryos are white to light yellow and oval, approximately 1.5 mm in length. Embryogenesis takes ~55 hr at 30 &#176;C. (B, C and G) Larvae have dark pigmented stripes and are covered with setae. Larvae go through several instars depending on the environment and their length can extend up to over 1 cm. (D, H) Young pupae are light yellow. Pupation takes ~ 5 - 7 days at 30 &#176;C. (E, I) Shortly after eclosion, dark pigmentation appears over the adult beetle body. Adults can live up to several months and one female can lay hundreds of embryos over her lifetime. <a href="http://cloudflare.jove.com/files/ftp_upload/54976/54976fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Previously, we showed that RNAi is effective in knocking down gene function in D. maculatus53. Here our experience rearing D. maculatus colonies in the laboratory is shared along with step-by-step protocols for both embryonic and parental RNAi set-up, injection, post-injection care, and phenotypic analysis. The dsRNA-mediated gene knockdown and analysis methods introduced here not only provide detailed information for addressing questions in D. maculatus, but also have potential significance for applying RNAi in other non-model beetle/insect species.

<table><tbody><tr><td>&lt;em&gt;Dermestes maculatus&lt;/em&gt; live beetles</td><td>Our lab or Carolina Biological Supply</td><td>#144168</td><td>Our lab strain was verified by COI barcoding; strain variation from Carolina cannot be ruled out</td></tr><tr><td>Wet cat food</td><td>Fancy Feast</td><td></td><td>Chunks of meat with gravy. Can buy at most pet food and grocery stores</td></tr><tr><td>Dry dog food</td><td>Purina Puppy Chow</td><td></td><td>Can buy at most pet food and grocery stores</td></tr><tr><td>Insect cage (size medium, 30.5 x 19 x 20.3 cm)</td><td>Exo Terra</td><td>PT2260</td><td>For colony maintenance. Can use larger cage if needed</td></tr><tr><td>Insect cage (size mini, 17.8 x 10.2 x 12.7 cm)</td><td>Exo Terra</td><td>PT2250</td><td>For embryo collection</td></tr><tr><td>Petri dish</td><td>VWR</td><td>89038-968</td><td></td></tr><tr><td>Cotton ball</td><td>Fisher</td><td>22-456-883</td><td></td></tr><tr><td>Megascript T7 transcription kit</td><td>Fisher</td><td>AM1334</td><td>For 40 reactions</td></tr><tr><td>Pneumatic pump</td><td>WPI</td><td>PV830</td><td></td></tr><tr><td>Capillary holder</td><td>WPI</td><td></td><td></td></tr><tr><td>Micromanipulator</td><td>NARISHIGE</td><td>MN-151</td><td></td></tr><tr><td>Black filter paper (90&amp;nbsp;mm)</td><td>VWR</td><td>28342-010</td><td></td></tr><tr><td>Food coloring (green)</td><td>McCormick</td><td></td><td></td></tr><tr><td>Borosilicate glass capillary</td><td>Hilgenberg</td><td>1406119</td><td></td></tr><tr><td>Needle puller (micropipette puller)</td><td>Sutter Instrument Co.</td><td>P-97</td><td></td></tr><tr><td>Microscope glass slide</td><td>WorldWide Life Sciences Division</td><td>41351157</td><td></td></tr><tr><td>Sealing film (Parafilm M)</td><td>Fisher</td><td>13-374-12</td><td></td></tr><tr><td>Model 801 Syringe (10&amp;nbsp;&amp;micro;l)</td><td>Hamilton</td><td>7642-01</td><td></td></tr><tr><td>Needle (32-gauge)&amp;nbsp;</td><td>Hamilton</td><td>7762-05</td><td></td></tr><tr><td>Fixation Solution (Pampel&#039;s)</td><td>BioQuip Products, Inc.</td><td>1184C</td><td>Toxic, needs to be handled in fume hood</td></tr><tr><td>Forcep (DUMONT #5)</td><td>Fine Science Tools</td><td>11252-30</td><td></td></tr><tr><td>Cover slip (24 x 50 mm, No. 1.5)</td><td>Globe Scientific</td><td>1415-15</td><td></td></tr><tr><td>Eppendorf Femtotips Microloader pipette tip</td><td>Fisher</td><td>E5242956003</td><td></td></tr><tr><td>Dissecting microscopy for embryo injection</td><td>Leica</td><td>M420</td><td></td></tr><tr><td>Dissecting microscopy for larval phenotypic visualization</td><td>Zeiss</td><td>SteREO Discover. V12</td><td></td></tr><tr><td>DIC microscopy</td><td>Zeiss</td><td>AXIO Imager. M1</td><td></td></tr></tbody></table>

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.

The authors&#39; lab has used RNAi technology to study the functional evolution of genes regulating segmentation in insects53,55. While all insects are segmented, the genes regulating this process appear to have diverged during insect radiations26,38,56-63. Genetic screens in Drosophila identified a set of nine pair-rule segmentation genes that are responsible for promoting the formation of body segments64-70. Here, the ortholog of one of these ge...

Watch this Scientific Journal Video about Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus at JoVE.com

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.

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 ...

rearing-double-stranded-rna-mediated-gene-knockdown-hide-beetle

Research

JoVE Journal

Genetics

10.6K Views.  University of Maryland. 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.

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

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 ...

Environment

RNAi-mediated Double Gene Knockdown and Gustatory Perception Measurement in Honey Bees (Apis mellifera)

This video demonstrates novel techniques of RNA interference (RNAi) which downregulate two genes simultaneously in honey bees using double-stranded RNA (dsRNA) injections. It also presents a protocol of proboscis extension response (PER) assay for measuring gustatory perception.	
	RNAi-mediated gene knockdown is an effective technique downregulating target gene expression. This technique is usually used for single gene manipulation, but it has limitations to detect interactions and joint effects between genes. In the first part of this video, we present two strategies to simultaneously knock down two genes (called double gene knockdown). We show both strategies are able to effectively suppress two genes, vitellogenin (vg) and ultraspiracle (usp), which are in a regulatory feedback loop. This double gene knockdown approach can be used to dissect interrelationships between genes and can be readily applied in different insect species.	
	The second part of this video is a demonstration of proboscis extension response (PER) assay in honey bees after the treatment of double gene knockdown. The PER assay is a standard test for measuring gustatory perception in honey bees, which is a key predictor for how fast a honey bee's behavioral maturation is. Greater gustatory perception of nest bees indicates increased behavioral development which is often associated with an earlier age at onset of foraging and foraging specialization in pollen. In addition, PER assay can be applied to identify metabolic states of satiation or hunger in honey bees. Finally, PER assay combined with pairing different odor stimuli for conditioning the bees is also widely used for learning and memory studies in honey bees.

RNAi-mediated Double Gene Knockdown and Gustatory Perception Measurement in Honey Bees Apis mellifera

In this protocol, we describe two strategies that simultaneously suppress two genes (double gene knockdown) in honey bees. Then we present how to use the proboscis extension response (PER) assay to study the effect of double gene knockdown on honey bee gustatory perception.

This video demonstrates novel techniques of RNA  interference (RNAi) which downregulate two genes simultaneously in honey bees  using double-stranded RNA ...

Neuroscience

Electroporation-mediated RNA Interference Method in Odonata

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

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

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

Developmental Biology

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

RNAi Mediated Gene Knockdown and Transgenesis by Microinjection in the Necromenic Nematode Pristionchus pacificus

Although it is increasingly affordable for emerging model organisms to obtain completely sequenced genomes, further in-depth gene function and expression analyses by RNA interference and stable transgenesis remain limited in many species due to the particular anatomy and molecular cellular biology of the organism. For example, outside of the crown group Caenorhabditis that includes Caenorhabditis elegans3, stably transmitted transgenic lines in non-Caenorhabditis species have not been reported in this specious phylum (Nematoda), with the exception of Strongyloides stercoralis4 and Pristionchus pacificus5. To facilitate the expanding role of P. pacificus in the study of development, evolution, and behavior6-7, we describe here the current methods to use microinjection for making transgenic animals and gene knock down by RNAi. Like the gonads of C. elegans and most other nematodes, the gonads of P. pacificus is syncitial and capable of incorporating DNA and RNA into the oocytes when delivered by direct microinjection. Unlike C. elegans however, stable transgene inheritance and somatic expression in P. pacificus requires the addition of self genomic DNA digested with endonucleases complementary to the ends of target transgenes and coinjection markers5. The addition of carrier genomic DNA is similar to the requirement for transgene expression in Strongyloides stercoralis4 and in the germ cells of C. elegans. However, it is not clear if the specific requirement for the animals' own genomic DNA is because P. pacificus soma is very efficient at silencing non-complex multi-copy genes or that extrachromosomal arrays in P. pacificus require genomic sequences for proper kinetochore assembly during mitosis. The ventral migration of the two-armed (didelphic) gonads in hermaphrodites further complicates the ability to inject both gonads in individual worms8. We also demonstrate the use of microinjection to knockdown a dominant mutant (roller,tu92) by injecting double-stranded RNA (dsRNA) into the gonads to obtain non-rolling F1 progeny. Unlike C. elegans, but like most other nematodes, P. pacificus PS312 is not receptive to systemic RNAi via feeding and soaking and therefore dsRNA must be administered by microinjection into the syncitial gonads. In this current study, we hope to describe the microinjection process needed to transform a Ppa-egl-4 promoter::GFP fusion reporter and knockdown a dominant roller prl-1 (tu92) mutant in a visually informative protocol.

RNAi Mediated Gene Knockdown and Transgenesis by Microinjection in the Necromenic Nematode Pristionchus pacificus

In model organisms, transgenesis can manipulate gene functions while RNAi can knockdown specific mRNA transcripts 1-2. This protocol aims to illustrate the techniques needed to introduce stably transmitted DNA and transient double stranded RNA into the necromenic nematode Pristionchus pacificus for studies in evolutionary, developmental, and behavioral biology.

Although it is increasingly affordable for emerging model organisms to obtain completely sequenced genomes, further in-depth gene function and expression ...

Large-scale Gene Knockdown in C. elegans Using dsRNA Feeding Libraries to Generate Robust Loss-of-function Phenotypes

RNA interference by feeding worms bacteria expressing dsRNAs has been a useful tool to assess gene function in C. elegans. While this strategy works well when a small number of genes are targeted for knockdown, large scale feeding screens show variable knockdown efficiencies, which limits their utility. We have deconstructed previously published RNAi knockdown protocols and found that the primary source of the reduced knockdown can be attributed to the loss of dsRNA-encoding plasmids from the bacteria fed to the animals. Based on these observations, we have developed a dsRNA feeding protocol that greatly reduces or eliminates plasmid loss to achieve efficient, high throughput knockdown. We demonstrate that this protocol will produce robust, reproducible knock down of C. elegans genes in multiple tissue types, including neurons, and will permit efficient knockdown in large scale screens. This protocol uses a commercially available dsRNA feeding library and describes all steps needed to duplicate the library and perform dsRNA screens. The protocol does not require the use of any sophisticated equipment, and can therefore be performed by any C. elegans lab.

Large-scale Gene Knockdown in C. elegans Using dsRNA Feeding Libraries to Generate Robust Loss-of-function Phenotypes

While dsRNA feeding in C. elegans is a powerful tool to assess gene function, current protocols for large scale feeding screens result in variable knockdown efficiencies. We describe an improved protocol for performing large scale RNAi feeding screens that results in highly efficacious and reproducible knockdown of gene expression.

RNA interference by feeding worms bacteria expressing dsRNAs has been a useful tool to assess gene function in C. elegans. While this strategy works well ...

Larval RNA Interference in the Red Flour Beetle, Tribolium castaneum

The red flour beetle, Tribolium&#160;castaneum, offers a repertoire of experimental tools for genetic and developmental studies, including a fully annotated genome sequence, transposon-based transgenesis, and effective RNA interference (RNAi). Among these advantages, RNAi-based gene knockdown techniques are at the core of Tribolium research. T. castaneum show a robust systemic RNAi response, making it possible to perform RNAi at any life stage by simply injecting double-stranded RNA (dsRNA) into the beetle&#8217;s body cavity.
In this report, we provide an overview of our larval RNAi technique in T. castaneum. The protocol includes (i) isolation of the proper stage of T. castaneum larvae for injection, (ii) preparation for the injection setting, and (iii) dsRNA injection. Larval RNAi is a simple, but powerful technique that provides us with quick access to loss-of-function phenotypes, including multiple gene knockdown phenotypes as well as a series of hypomorphic phenotypes. Since virtually all T. castaneum tissues are susceptible to extracellular dsRNA, the larval RNAi technique allows researchers to study a wide variety of tissues in diverse contexts, including the genetic basis of organismal responses to the outside environment. In addition, the simplicity of this technique stimulates more student involvement in research, making T. castaneum an ideal genetic system for use in a classroom setting.

Larval RNA Interference in the Red Flour Beetle, Tribolium castaneum

RNA interference (RNAi)-based gene knockdown techniques are at the core of Tribolium research. Here, we provide an overview of our larval RNAi technique in Tribolium castaneum. Larval RNAi is a simple, but powerful technique that provides quick access to loss-of-function phenotypes, allowing researchers to study gene functions in diverse contexts.

The red flour beetle, Tribolium&#160;castaneum, offers a repertoire of experimental tools for genetic and developmental studies, including a fully ...

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 ...

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

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

RNAi-mediated Double Gene Knockdown and Gustatory Perception Measurement in Honey Bees (Apis mellifera)

Electroporation-mediated RNA Interference Method in Odonata

Applications of RNA Interference in American Cockroach

RNAi Interference by dsRNA Injection into Drosophila Embryos

RNAi Mediated Gene Knockdown and Transgenesis by Microinjection in the Necromenic Nematode Pristionchus pacificus

Large-scale Gene Knockdown in C. elegans Using dsRNA Feeding Libraries to Generate Robust Loss-of-function Phenotypes

Larval RNA Interference in the Red Flour Beetle, Tribolium castaneum

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

RNAi Trigger Delivery into Anopheles gambiae Pupae