We would like to thank Dr. Flaminia Catteruccia and her research group (Harvard School of Public Health, Boston, MA) for providing our laboratory with the G3 colony used in this research and for the LacZ template DNA. We thank Adam Jenkins (Boston College, Chestnut Hill, MA) for his assistance in maintaining the insectary. The Biology Department of Boston College generously funded the research associated with the development of this technique.

1. Synthesis and Preparation dsRNA
<ol>
	<li>Identify a 200-800 bp knockdown region (to generate the corresponding dsRNA) within the gene of interest that is predicted to have no identifiable off-target effects (e.g., no sequence homology &#8805;18 bp within another gene) and a negative control (e.g., a heterologous sequence that is not present within target insect genome, such as the Escherichia colilacZ gene (GenBank Gene ID: 945006). Alternatively, use a positive control (e.g., which yields an easily observed phenotype, such as SRPN221,22, GenBank Gene ID: 1270169). The sequence of the dsRNA targeting SRPN2 is defined in Michel et al. (2005)21. 
	Note: E-RNAi is an open-source bioinformatic resource that is useful for the identification of such regions and for the process of designing oligonucleotide primers23.</li>
	<li>Perform standard PCR amplification (i.e., performed with Taq DNA polymerase using 30-35 cycles) using a genomic DNA or cDNA template to obtain dsRNA synthesis template flanked by a T7 promoter sequence (5&#39;-TAATACGACTCACTATAGGG-3&#39;) and proceed with dsRNA preparation and clean-up using a commercial in vitro transcription kit as per manufacturer&#39;s instructions. Use SRPN2 PCR amplification conditions and primer information as presented in An et al. (2011)22.</li>
	<li>Quantify purified RNA amplicon yields by ultraviolet absorbance spectroscopy at wavelength of 260 nm and adjust to the desired concentration (e.g., 3 &#181;g/&#181;l) in RNase-free water.
	<ol>
		<li>For troubleshooting low RNA concentrations, reduce liquid volume by spinning samples down in a vacuum centrifuge at room temperature or by lyophilizing samples and reconstituting in smaller volumes of water. The time required for sample lyophilization will vary depending on initial sample volumes and dsRNA concentrations.</li>
	</ol>
	</li>
	<li>Check the quality and length of the dsRNA on a 2% agarose gel prepared with 1x TBE or TAE buffer and stained with ethidium bromide (EtBr), along with the template DNA used for the transcription reaction. The dsRNA will migrate more slowly than template DNA. Quality and length can be assessed by assuring there are no non-specific dsRNA products and by comparing products with a standard DNA marker, respectively. 
	Note: EtBr is a potent mutagen and should be handled accordingly. 
	Note: At a dsRNA concentration of 3 &#181;g/&#181;l, 0.5 &#181;l of the sample is more than sufficient for visualization on the EtBr-stained agarose gel.</li>
	<li>Store dsRNA at -20 &#176;C until needed. Multiple freeze/thaw cycles may cause degradation, so aliquots should be prepared for large volumes of dsRNA.</li>
</ol>
2. Prepare Fast Green FCF Dye (FGD) Tubes
<ol>
	<li>Dilute Fast Green FCF dye from stock solution (&#8805;85% dye content) to 0.1% (v/v) (working solution) in RNase-free water.</li>
	<li>Pipette 1 &#181;l of dye into the bottom of a 1.5 ml microcentrifuge tube.</li>
	<li>Place tubes in a 65 &#176;C heat block for approximately 3 hr to evaporate liquid, then place tubes at room temperature for at least 30 min, to cool before using. This dry solid dye will reconstitute in dsRNA resuspension solution.</li>
</ol>
3. Pull Injection Needles
<ol>
	<li>Pull borosilicate glass needles using a heated needle puller to a tip diameter of 10-30 &#181;m. Pull settings correspond to: Heater adjustment no. 1 = 100, Heater adjustment no. 2 = 70.</li>
	<li>To avoid damage to the fine tip of the needle, place all pulled needles horizontally in a Petri dish on a strip of molding putty. 
	&#8203;Note: Additional information for capillary needle pulling methods can be found in further detail in the Malaria Research and Reference Reagent Resource Center MR4 manual24.</li>
</ol>
4. Prepare Injection Station
<ol>
	<li>Collect materials required: glass microcapillary needles pulled to fine tip, microinjector, thin filter paper and thick filter paper, Petri dishes, transfer pipettes, paint brush and dissecting light microscope.</li>
	<li>Prepare the microinjector as instructed in the microinjector manual, and set injection volume to desired volume per pulse (e.g., maximum of 69 nl per pulse).</li>
	<li>On a platform that is easy to maneuver under a microscope (e.g., flat side of a Styrofoam tube rack), stack the two filter paper sheets with the thin filter paper on top, and secure with tape around the edges.</li>
	<li>Resuspend 10 &#181;l of each dsRNA solution in separate colored dye tubes, and place on ice.</li>
</ol>
5. Collect Pupae for Injection
<ol>
	<li>Fill a small 60 mm x 15 mm Petri dish with 10 ml of deionized water, and collect ~50 pale pupae (during the first 24 hr after pupation) from an insectary tray using a disposable plastic transfer pipette.</li>
	<li>Remove any pupae that have medium to dark cuticle tanning. 
	&#8203;Note: Once the cuticle begins to tan, it becomes more difficult to penetrate the cuticle, and injection results in much higher lethality.</li>
</ol>
6. dsRNA Injection
<ol>
	<li>Under the dissecting microscope, break off the distal tip of the injection needle with a pair of fine forceps.</li>
	<li>Prepare the injection needle by backfilling the needle with mineral oil (using a syringe with a 3 inch, 30 gauge needle), and expelling excess oil with the microinjector.</li>
	<li>Front fill injection needle with maximum amount of dsRNA, and eject one pulse under the microscope to ensure the dispensing of liquid. In the event that no liquid is taken up and/or expelled, check the distal tip of the needle for any blockage and ensure that the needle is firmly secured in the microinjector.</li>
	<li>Pick 1-3 pupae, and place them onto the filter paper.</li>
	<li>Using the paintbrush, position the pupae on the filter paper with dorsal cuticle accessible, and use the paintbrush to push on filter paper and absorb of any excess water.</li>
	<li>Stabilize the pupa with the tip of the paintbrush, and insert the needle into the dorsal cuticle between the thorax and abdomen at an angle of approximately 30&#176; in relation to the dorsal surface of the pupa. Injection should be directed toward the posterior end of the pupa and needle should be moved in an anterior-to-posterior direction.</li>
	<li>Inject two pulses (69 nl per pulse) of 3 &#181;g/&#181;l dsRNA solution into the hemolymph, and check for the distribution of color throughout the body. If no color is identified, shift the injection needle position slightly to clear the tip from obstruction and repeat liquid delivery.</li>
	<li>Use the wetted paintbrush to gently move pupa from the needle into water for culturing. The pupa should stick to the paintbrush upon light contact.</li>
</ol>
7. Post-injection Conditions
<ol>
	<li>Place Petri dish with injected pupae into a mosquito cage with suitable airflow (e.g., mesh cage or container with mesh lid). Pupal rearing conditions should remain consistently at 27 &#177; 3 &#176;C temperature, 75 &#177; 5% humidity and be under a light:dark cycle of 16:8 hr.</li>
	<li>Prepare a 10% (w/v) glucose solution, and place a solution-saturated cotton ball on the mosquito cage mesh for adult feeding.</li>
</ol>
8. Assess Knockdown
<ol>
	<li>At desired time-point(s), assess phenotypes in experimental dsRNA-injected insects, compared to controls.
	<ol>
		<li>Assess dsSRPN2 and dsLacZ insects daily by chilling down adults for about 2-3 min at -20 &#176;C, transferring them to a cold plate at 2 &#176;C and identifying any pseudo-tumor formation by utilization of a dissecting microscope at 15X magnification with brightfield illumination. After assessment, return adults to insectary conditions (27 &#177; 3 &#176;C, 75 &#177; 5% humidity). 
		Note: The experimental and control dsRNAs employed in this protocol are dsSRPN2 and dsLacZ, respectively. There are many options suitable for controls; however, it is suggested that a positive control for which phenotype and/or expression is easily visualized (e.g., by dissecting microscopy) and/or quantified [e.g., quantitative real-time PCR (qRT-PCR), Western blot] should be used when learning this technique. SRPN2 protein and transcript quantification via Western blot and qRT-PCR, respectively, are described in Michel et al. (2005)21.</li>
	</ol>
	</li>
</ol>

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H., Meyering-Vos, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+interference+with+the+allatoregulating+neuropeptide+genes+from+the+fall+armyworm+Spodoptera+frugiperda+and+its+effects+on+the+JH+titer+in+the+hemolymph.">RNA interference with the allatoregulating neuropeptide genes from the fall armyworm Spodoptera frugiperda and its effects on the JH titer in the hemolymph.</a> J. Insect Physiol. 54 (6), 997-1007 (2008).</li><li>Scott, J. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Towards+the+elements+of+successful+insect+RNAi.">Towards the elements of successful insect RNAi.</a> J. Insect Physiol. 59 (12), 1212-1221 (2013).</li></ol>

Open Access for this video-article is paid for by Biogen.

Current methods for inducing non-transgenic RNAi in mosquitoes involve direct injection of dsRNA into the adult hemocoel12,13 or larval feeding of RNAi trigger-coated nanoparticles14-17 or microalgae-based dsRNA molecules18. Targeting the adult mosquito, while extremely valuable, can exclude a large number of genes that function during earlier developmental periods. Knockdown initiated by larval feeding may yield inconsistent phenotypes during the adult stage due, in part, to the potential of variable protein persistence through the pupal stage. Therefore, introducing an additional method that is aimed specifically at initiating RNAi during pupal development will provide a means to more fully assess gene functions during pre-adult developmental stages, as well as enhanced abilities to assess gene function during adult stages. As with gene knockdown approach based on dsRNA injection or expression, the persistence of gene knockdown cannot be predicted. Therefore, transcript or protein levels should be assessed for gene of interest during developmental periods of interest. Although we observe a continuation of decreased protein levels at day 5 post-injection for SRPN2 in SRPN2 dsRNA-injected animals, factors such as protein turnover and half-life can differ for different targets. It is critical to ensure, as well, that the dsRNA to be used for injection is well concentrated and appears intact on an agarose gel. We recommend experimenters reassess these factors if knockdown results are not sufficient, and respective concentrations of dsRNA may need to be tested empirically for specific gene targets.
We describe a method for the initiation of RNA interference during the pupal stage of An. gambiae development. This method relies on the introduction via microinjection of dsRNA directly into the hemocoel of early pupae and allows for assessment of injection quality by the use of dye-labeled dsRNA. The ability to visualize injection quality constitutes a critical enhancement for ensuring successful knockdown and constitutes an aspect of injection-based gene knockdown that has not been considered in most previously reported dsRNA-based protocols focusing on the adult stage. By targeting the pupa at the onset of this developmental period, genes that might play a role during this critical developmental interval, or during the early stages of adulthood can be evaluated functionally. Additionally, this method may enable dsRNA delivery to cells, and establishment of RNA interference in cells that are accessible during metamorphosis, but less accessible in fully formed adult mosquitoes.
A recent microarray analysis by Harker et al. (2012) identified 560 An. gambiae transcripts that were up-regulated or down-regulated by at least 4-fold during distinct developmental stages, ranging from the embryo to adult. Of the 560 transcripts identified, a set of 309 was up-regulated during pupal development27. These findings suggest that there are many requirements for differential gene expression throughout mosquito development, including those that occur during the pupal stage, an interval during which the organism undergoes metamorphosis. In many insect species, including An. gambiae, genes involved in processes such as development (i.e., pupal cuticular and chitin-binding proteins)27-31 and immune response (i.e., Toll receptor-like proteins)27,32-34 are highly expressed during the pupal stage. Once a fully formed adult has emerged, there is continued gene expression in response to environmental and physiological changes35. Notably, during early adult development, there is an increase in the expression of developmental genes (i.e., adult cuticular and sarcoplasmic proteins)36, as well as other key genes (i.e., sperm specific protein and cytochrome P450 metabolism enzymes)27,36.
The positive control used in the development of this protocol, SRPN2, is an An. gambiae serine protease inhibitor (serpin). SRPN2 plays an important role in the negative regulation of insect melanization, a broad spectrum innate immune response in insects21,22. Knockdown of SRPN2 in adult mosquitoes results in pseudo-tumor formation21,22, a phenotype that is easily observed by use of light microscopy. Given that this distinct phenotype can be easily scored in live insects, we used SRPN2 for initial pupal stage RNAi injections. In addition, SRPN2 is expressed during all developmental stages37, thereby providing a good target for pupal stage RNAi injection and assessment of function in the early adult. We demonstrate that the method we have developed is capable of inducing similar adult melanotic pseudo-tumor formation as a consequence of dsRNA injections performed during the pupal stage of development. In developing this protocol, we have observed that injection during early pupal development (i.e., the first 24 hr after the larval-pupal molt) is critical for obtaining optimal adult emergence. In the event that poor emergence is obtained post-injection, we recommend staging larvae with greater accuracy, to obtain pupae with less extensive cuticle hardening and assure early pupal stage injection is achieved. Furthermore, minimizing damage to the cuticle results in optimal emergence rates and increasing magnification during injection can help ensure that only the intended region of the animal is punctured. If the needle should puncture through to the ventral cuticle, pooling of dye-labeled liquid will be evident on the exterior of the pupa or will saturate the filter paper. It is highly recommended to discard any pupae that have more than one cuticle puncture.
With the extensive experiences of many laboratories with the performance of adult mosquito injections, previously identified microinjection approaches can be adapted with simple protocol modifications for use in pupal RNAi experiments. Overall, the goal of this method is to provide researchers the ability to expand the timeframe during which reverse genetic analyses can be performed, further enabling research that will support the development of novel vector control strategies. Interestingly, experiments in other species, such as Rhodnius prolixus and Spodoptera frugiperda, reveal that gene silencing effects tend to be much greater when initiated during pre-adult stages38,39. During all stages of development, RNAi-mediated gene knockdown is subject to considerations regarding the rapidity and persistence of gene silencing, and the stability of proteins encoded by targeted genes. The ideal RNAi target genes tend to be those that encode a protein or RNA that has a short half-life and high turnover rate11,40.
While transgenic RNAi strategies can also be employed to address considerations regarding rapidity and persistence of RNAi during pre-adult stages, transgenic techniques have many drawbacks (e.g., time required for the generation of transgenic lines, experimental time-frames for mosquito matings to generate insects with regulated dsRNA expression, and maintenance of transgenic stocks). By contrast, our protocol affords an easier and faster method for initiating gene knockdown during pupal development and in cell types that originate and are accessible during metamorphosis but are less accessible in adults. The use of dye-labeled dsRNA suspensions allows for easy assessment of injection success and dispersal of introduced material within pupae. Our data on the onset of tumor formation in pupal-injected adults, as compared to animals injected as adults, is consistent with initiation of RNAi-mediated knockdown during the pupal stage. Using our G3 mosquito line and under our insectary conditions, we observe the adult melanotic pseudo-tumor formation as early as 10 days post-injection of adults injected three to five days post-emergence. By performing early pupal-stage injection, we observe visible melanotic pseudo-tumor formation as early as five days post-injection (i.e., three to four days post emergence). These data imply that this method enables initiation of gene knockdown during a previously under-studied developmental period (i.e., pupal development). Our dye labeling method may also prove useful for the development of new larval injection protocols, due to the translucent nature of the cuticle during all larval instars. While the control used in this study requires progression into adult stage to display a knockdown phenotype, future experiments to assess pupal stage-specific phenotypes following injection of early pupae will provide valuable insight regarding the expansion of this method into additional developmental periods such as late pupal development. In summary, this method provides a valuable RNAi protocol for pupal stage initiation of RNAi-mediated gene knockdown and expands the functional genomic tools available for use within the vector insect research community.

Malaria is a mosquito-borne infectious disease that affects many millions of individuals every year. The World Health Organization (WHO) reports that in 2013 there were approximately 584,000 deaths due to malaria, 78 percent of which occurred in children under the age of five years1. The pathogens that cause human malaria are apicomplexan parasites within the genus Plasmodium and are transmitted between their human hosts by female Anopheles mosquitoes. Transmission occurs when the mosquito takes a blood meal from an individual who is infected, and then deposits infective parasites into an uninfected individual in a subsequent blood meal. Within the genus Anopheles, Anopheles gambiae is the species with the greatest vectorial capacity and is the most prominent malaria vector in sub-Saharan Africa1-3.
Currently, mosquito vector control by deployment of insecticides continues to be the major method employed to reduce the burden of human malaria. Although the use of insecticides since the 1960s has proven to be extremely successful, the rise of insecticide resistance has driven a need for development of novel insecticides and alternative vector control strategies4-7. During 2010, a total of 49 of 63 countries reporting to the WHO indicated the occurrence of insecticide resistance in malaria vectors1. Additionally, the IR Mapper tool, which utilizes peer-reviewed literature to assess resistance data in Afrotropical regions, reports that between 2001 and 2012 there were 46% and 27% increases in resistance to pyrethroids and dichlorodiphenyltrichloroethane (DDT), respectively7.
RNA interference (RNAi) was identified in the early 1990s as a technique that could be employed to inactivate genes in the Petunia plant8,9 and in the fungus Neurospora crassa9,10. Shortly thereafter, in 1998, RNAi was first documented in Caenorhabditis elegans as a means of reducing gene expression in an animal model by introduction of antisense or double-strand RNA (dsRNA) via injection or feeding methods9,11. Since its discovery, RNAi has revolutionized the pursuit of functional genomics by allowing researchers to utilize reverse genetics to rapidly investigate the functional roles of genes of interest via a highly selective post-transcriptional gene silencing mechanism. In some organisms, such as Drosophila melanogaster, the use of transgenic organisms that express interfering RNA constructs has been widely successful for gene knockdown (KD). Although the use of transgenes in An. gambiae for RNAi has been utilized and may prove useful for large-scale screens, the generation of transgenic mosquito strains is both labor and time intensive, generally taking two to three months to go from the identification of a gene of interest to the generation of an appropriate transgenic stock12. Currently, the primary method of gene KD in An. gambiae is by injection into the hemolymph, during the adult stage, of dsRNA specific for a given gene12,13. This process typically takes about one month to go from identification of a gene of interest to assessment of gene KD, proving to be much more rapid than transgenic methods12. Methods for larval-stage RNAi have been established recently in An. gambiae and Aedes aegypti via nanoparticle feeding14-17 or by oral delivery of microalgae-based dsRNA molecules18, offering opportunities to perform functional genomic analysis during early stages of development. In direct injection, feeding, and nanoparticle delivery methods, dsRNA is taken up autonomously by the target cell and cleaved by the enzyme Dicer into 21-25 nucleotide-long &#34;short interfering RNAs&#34; (siRNAs)19,20. These siRNAs are then incorporated into the RNA-induced silencing complex (RISC), from which one strand will be discarded, allowing the RNA-bound RISC complex to bind to and cleave the target mRNA and thereby reduce its level and inhibit its translation19,20.
Many intrinsic features of basic mosquito biology modulate vectorial capacity, including host preference (e.g., olfaction, gustation), mating, reproduction and immunity. Given the importance of these biological processes, it is likely that their modulation on a genetic or pharmacological level will offer new opportunities for vector control, including circumvention of insecticide resistance, and provide new tools for more broadly integrated approaches to vector management. The use of functional genomics to assess the roles of genes underlying these intrinsic biological features will enable identification of new targets and provide new insights into how we can effectively create new, more effective control strategies. We describe the development and use of a rapid injection method for initiating RNAi during the pupal stage of An. gambiae. We observe that pupal stage injection of an RNAi trigger enables observation of resultant phenotypes in early stage adults, i.e., sooner after emergence than would be observed if gene knockdown were initiated in adults post-emergence. This methods enables gene knockdown beginning during the pupal developmental interval and extending into adult stages, such that gene knockdown initiated during pupal development can persist and affect early adult hemolymph-accessible cell types, as well as cell types that are more hemolymph-accessible during metamorphosis than in the adult, such as sensory neurons found in adult appendages following emergence.

<table>
	<tbody>
		<tr>
			<td>MEGAscript RNAi kit</td>
			<td>Ambion</td>
			<td>AM1626</td>
			<td></td>
		</tr>
		<tr>
			<td>Nanoject II injector</td>
			<td>Drummond</td>
			<td>3-000-204</td>
			<td></td>
		</tr>
		<tr>
			<td>Nanoject II foot switch</td>
			<td>Drummond</td>
			<td>3-000-026</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate glass capillaries</td>
			<td>Drummond</td>
			<td>3-000-203-G/X</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass micropipette puller</td>
			<td>Narishigne</td>
			<td>PB-7</td>
			<td></td>
		</tr>
		<tr>
			<td>Fast Green FCF dye</td>
			<td>Sigma</td>
			<td>F7258</td>
			<td>Can substitute with a non-toxic food dye.</td>
		</tr>
		<tr>
			<td>Plastic transfer pipettes</td>
			<td>Thermo Scientific</td>
			<td>1371150</td>
			<td>&#188; inch cut from tip to create a wider opening.</td>
		</tr>
		<tr>
			<td>Whatman &#8220;thin&#8221; filter paper&#160;</td>
			<td>GE Healthcare</td>
			<td>1001-090</td>
			<td>This is 9 cm diameter Grade 1, but can be altered depending on size of injection pad.</td>
		</tr>
		<tr>
			<td>Western blot &#8220;thick&#8221; filter paper</td>
			<td>BioRad</td>
			<td>107-3931</td>
			<td>This is the filter paper generally used for Western blots.</td>
		</tr>
		<tr>
			<td>Petri dishes (60 mm x 15 mm)</td>
			<td>Fisher Scientific</td>
			<td>FB0875713A</td>
			<td></td>
		</tr>
		<tr>
			<td>Paint brush (size 0)</td>
			<td>Michael&#8217;s Art</td>
			<td>10474940</td>
			<td>Brush size and brand can vary based on availability and user preference.</td>
		</tr>
		<tr>
			<td>Dissecting light microscope</td>
			<td>&#160;----&#160;</td>
			<td>-----</td>
			<td>This can vary based on availability and user preference.</td>
		</tr>
	</tbody>
</table>

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.

Pupal injection for gene KD yields optimal results when injection is performed during the early pupal stage, when cuticle tanning levels are low (Figure 1A, left, and 1B). Increased tanning and hardening of cuticle, generally after 24 hours, results in increased pupal death following injection (Figure 1A, center and right). The rate of pupal development can vary depending on insectary conditions and animal density24,25; therefore, it is best to assess pigmentation visually.
During the injection process, the capillary needle is inserted into the dorsal cuticle at an angle of approximately 30&#176; in the anterior to posterior direction (Figure 2A). Once the needle is inserted and the dsRNA + 0.01% (w/v) FGD is dispensed, the distribution of dye is evident throughout the hemolymph (Figure 2B).
Assessment of adult emergence for pupae injected with 0.01% (w/v) FGD revealed an average rate of 70% emergence, compared to 96.7% emergence of non-injected controls (Figure 3A). Of note, partial emergence from the pupal case was observed for a large number of non-surviving mosquitoes (Figure 3B). Injected animals exhibit no delays in emergence time (Figure 4A) or biased impact on either gender (Figure 4B). Additional assessment of adult survival carried out up to day 10 post-emergence reveals no evident impact on post-emergence adult survival (Figure 4C).
Validation of KD quality was assessed by the melanotic pseudo-tumor (darkly pigmented tissue cluster) phenotype associated with SRPN2 knockdown21,22 as a positive control for knockdown and the absence of phenotypes associated with dsLacZ injection as a negative control. Adult mosquitoes that emerged were scored at day 8 post-injection (day 6-7 post-emergence). Melanotic pseudo-tumors (Figure 5A and 5B) were observed through the cuticle of 93.5% of the dsSRPN2 vs. 0% of the dsLacZ adult mosquitoes (Figure 6A). Clusters of darkly melanized tissue were identified upon dissection of pigmented patches (Figure 5C). Pseudo-tumors were visible on the adult cuticle as early as day 3 post-emergence and were also present in a subset of dsSRPN2 hemolymph and gut tissues (data not shown). At 5 post-injection (early adult stage), significantly decreased SRPN2 levels in dsSRPN2, but not dsLacZ or non-injected hemolymph protein isolates was observed (Figure 6B).
<img alt="Figure 1" src="/files/ftp_upload/53738/53738fig1.jpg" /> 
Figure 1: Developmental staging for pupal dsRNA injection. Early pupal injection of dsRNA results in optimal survival and progression into adult stage. Low levels of cuticle pigmentation (A, left, and B) can be observed within the first 0-24 hr following pupation. Tanning of the pupal cuticle preceding injection (A, center and right) results in moderate to poor survival.
<img alt="Figure 2" src="/files/ftp_upload/53738/53738fig2.jpg" /> 
Figure 2: Injection position and distribution of dye-labeled dsRNA. (A) Capillary needle injection of dye-labeled dsRNA into the dorsal cuticle at an angle of approximately 30&#176;, in anterior to posterior direction. (B) The dye is visibly distributed in the pupal hemolymph. dsRNA injection volume of 138 nl, labeled with 0.01% FGD (w/v).
<img alt="Figure 3" src="/files/ftp_upload/53738/53738fig3.jpg" /> 
Figure 3: Post-injection adult emergence. (A) 70% of pupae injected with 0.01% FGD (w/v) successfully emerged (n = 60), compared to 96.7% of non-injected controls (n = 60). Three biological replicates were performed. (B) Partial emergence from the pupal case was observed for a large number of non-surviving mosquitoes. Error bars represent the standard error of the mean (SEM).
<img alt="Figure 4" src="/files/ftp_upload/53738/53738fig4.jpg" /> 
Figure 4: Emergence rate, sex assessment and adult survival. (A) Comparable emergence times were observed following pupal injection with 0.01% FGD (24 hr = 80% and 48 hr = 20%), as compared to non-injected pupae (24 hr = 83% and 48 hr = 17%). (B) Approximately equal male and female adult emergence was observed for 0.01% FGD injected pupae (female = 48% and male = 52%) and non-injected pupae (female = 52% and male = 48%). (C) Survival analysis reveals that injection with 0.01% FGD does not impact adult survival, assessed up to day 10 post-emergence. Results represent data from three independent experiments with 0.01% FGD injected (n = 60) and non-injected (n = 60) pupae (equal numbers of males and females). Error bars represent the standard error of the mean (SEM).
<img alt="Figure 5" src="/files/ftp_upload/53738/53738fig5.jpg" /> 
Figure 5: Pseudo-tumor positive control phenotype reflects successful knockdown. Pseudo-tumors were observed on the (A) abdominal and (B) thoracic cuticle of dsSRPN2-injected, but not dsLacZ-injected&#160;adult mosquitos at day 8 post-injection. (C) Higher magnification (400X) imaging (a) of cuticle and dissection of pigmented patches (b) reveals clusters of darkly melanized cells.
<img alt="Figure 6" src="/files/ftp_upload/53738/53738fig6.jpg" /> 
Figure 6: Quantification of pseudo-tumor formation and decreased SRPN2 protein levels. (A) Pupal stage injections result in pseudo-tumor formation in 93.5% of dsSRPN2 adults (n = 21) compared to 0% of dsLacZ controls (n = 19). Results obtained day 8 post-injection. (B) Western blot (left) shows decreased SRPN2 levels in dsSRPN2, but not dsLacZ or non-injected hemolymph protein isolates (day 5 post-injection). Results based on three independent experiments. Anti-SRPN221 and anti-SRPN326 antibody dilutions used were 1:1,000 and 1:2,000, respectively. Goat anti-rabbit IgG-HRP (Product sc-2004, Santa Cruz Biotechnology, Dallas TX) was used at 1:5,000. All protein levels were quantified (right) by band intensity (ImageJ Software, NIH, Bethesda, MD), normalized to SRPN3, and statistically compared by unpaired t test. P &#60;0.05: *, P &#8805;0.05: n.s. (not significant). Error bars represent the standard error of the mean (SEM).

Watch this Scientific Journal Video about RNAi Trigger Delivery into Anopheles gambiae Pupae at JoVE.com

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.

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 ...

rnai-trigger-delivery-into-anopheles-gambiae-pupae

Research

JoVE Journal

Biology

9.0K Views.  Boston College. 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.

Video: RNAi Trigger Delivery into Anopheles gambiae Pupae

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

Protocol for Mosquito Rearing (A. gambiae)

This protocol describes mosquito rearing in the insectary. The insectary rooms are maintained at 28&deg;C and ~80% humidity, with a 12 hr. day/night cycle. For this procedure, you'll need mosquito cages, 10% sterile sucrose solution, paper towels, beaker, whatman filter paper, glass feeders, human blood and serum, water bath, parafilm, distilled water, clean plastic trays, mosquito food (described below), mosquito net to cover the trays, vacuum, and a collection chamber to collect adults.

Protocol for Mosquito Rearing A. gambiae

This video illustrates the general techniques used to rear Anopheles gambiae in the laboratory. The methods for caring for laboratory mosquitoes are demonstrated through all stages of the organism's life cycle from larvae to pupae to blood-feeding adults.

This protocol describes mosquito rearing in the insectary. The insectary rooms are maintained at 28&deg;C and ~80% humidity, with a 12 hr. day/night ...

Protocol for RNAi Assays in Adult Mosquitoes (A. gambiae)

Reverse genetic approaches have proven extremely useful for determining  which genes underly resistance to vector pathogens in mosquitoes.  This video protocol illustrates a method used by the Dimopoulos lab to inject dsRNA into Anopheles gambiae mosquitoes, which harbor the malaria parasite.  The technique manipulating the injection setup and injecting dsRNA into the thorax is illustrated.

Protocol for RNAi Assays in Adult Mosquitoes A. gambiae

Reverse genetic approaches have proven extremely useful for determining which genes underly resistance to vector pathogens in mosquitoes. This video protocol illustrates a method used by the Dimopoulos lab to inject dsRNA into Anopheles gambiae mosquitoes, which harbor the malaria parasite. The technique manipulating the injection setup and injecting dsRNA into the thorax is illustrated.

Reverse genetic approaches have proven extremely useful for determining  which genes underly resistance to vector pathogens in mosquitoes.  This video ...

Localized RNAi and Ectopic Gene Expression in the Medicinal Leech

In this video, we show the use of a pneumatic capillary gun for the accurate biolistic delivery of reagents into live tissue. We use the procedure to perturb gene expression patterns in selected segments of leech embryos, leaving the untreated segments as internal controls.
The pneumatic capillary gun can be used to reach internal layers of cells at early stages of development without opening the specimen. As a method for localized introduction of substances into living tissues, the biolistic delivery with the gun has several advantages: it is fast, contact-free and non-destructive. In addition, a single capillary gun can be used for independent delivery of different substances. The delivery region can have lateral dimensions of ~50-150 &#181;m and extends over ~15 &#181;m around the mean penetration depth, which is adjustable between 0 and 50 &#181;m. This delivery has the advantage of being able to target a limited number of cells in a selected location intermediate between single cell knock down by microinjection and systemic knockdown through extracellular injections or by means of genetic approaches.
For knocking down or knocking in the expression of the axon guidance molecule Netrin, which is naturally expressed by some central neurons and in the ventral body wall, but not the dorsal domain, we deliver molecules of dsRNA or plasmid-DNA into the body wall and central ganglia. This procedure includes the following steps: (i) preparation of the experimental setup for a specific assay (adjusting the accelerating pressure), (ii) coating the particles with molecules of dsRNA or DNA, (iii) loading the coated particles into the gun, up to two reagents in one assay, (iv) preparing the animals for the particle delivery, (v) delivery of coated particles into the target tissue (body wall or ganglia), and (vi) processing the embryos (immunostaining, immunohistochemistry and neuronal labeling) to visualize the results, usually 2 to 3 days after the delivery.
When the particles were coated with netrin dsRNA, they caused clearly visible knock-down of netrin expression that only occurred in cells containing particles (usually, 1-2 particles per cell). Particles coated with a plasmid encoding EGFP induced fluorescence in neuronal cells when they stopped in their nuclei.

In this video, we show a procedure for an accurate biolistic delivery of reagents into live tissue with a novel miniature gene gun. We are knocking down the expression of the axon guidance molecule Netrin in leech embryos by delivering molecules of dsRNA into the ventral body wall and ganglia of single segments.

In this video, we show the use of a pneumatic capillary gun for the accurate biolistic delivery of reagents into live tissue. We use the procedure to ...

MISSION esiRNA for RNAi Screening in Mammalian Cells

RNA interference (RNAi) is a basic cellular mechanism for the control of gene expression. RNAi is induced by short double-stranded RNAs also known as small interfering RNAs (siRNAs). The short double-stranded RNAs originate from longer double stranded precursors by the activity of Dicer, a protein of the RNase III family of endonucleases. The resulting fragments are components of the RNA-induced silencing complex (RISC), directing it to the cognate target mRNA. RISC cleaves the target mRNA thereby reducing the expression of the encoded protein1,2,3. RNAi has become a powerful and widely used experimental method for loss of gene function studies in mammalian cells utilizing small interfering RNAs.
Currently two main methods are available for the production of small interfering RNAs. One method involves chemical synthesis, whereas an alternative method employs endonucleolytic cleavage of target specific long double-stranded RNAs by RNase III in vitro. Thereby, a diverse pool of siRNA-like oligonucleotides is produced which is also known as endoribonuclease-prepared siRNA or esiRNA. A comparison of efficacy of chemically derived siRNAs and esiRNAs shows that both triggers are potent in target-gene silencing. Differences can, however, be seen when comparing specificity. Many single chemically synthesized siRNAs produce prominent off-target effects, whereas the complex mixture inherent in esiRNAs leads to a more specific knockdown10.
In this study, we present the design of genome-scale MISSION esiRNA libraries and its utilization for RNAi screening exemplified by a DNA-content screen for the identification of genes involved in cell cycle progression. We show how to optimize the transfection protocol and the assay for screening in high throughput. We also demonstrate how large data-sets can be evaluated statistically and present methods to validate primary hits. Finally, we give potential starting points for further functional characterizations of validated hits.

Here we use a human esiRNA library in a high-throughput screen for genes involved in cell division. We demonstrate how to set up and conduct an esiRNA screens, as well as how to analyze and validate the results.

RNA interference (RNAi) is a basic cellular mechanism for the control of gene expression. RNAi is induced by short double-stranded RNAs also known as ...

Bacterial Delivery of RNAi Effectors: Transkingdom RNAi

RNA interference (RNAi) represents a high effective mechanism for specific inhibition of mRNA expression. Besides its potential as a powerful laboratory tool, the RNAi pathway appears to be promising for therapeutic utilization. For development of RNA interference (RNAi)-based therapies, delivery of RNAi-mediating agents to target cells is one of the major obstacles. A novel strategy to overcome this hurdle is transkingdom RNAi (tkRNAi). This technology uses non-pathogenic bacteria, e.g. Escherichia coli, to produce and deliver therapeutic short hairpin RNA (shRNA) into target cells to induce RNAi. A first-generation tkRNAi-mediating vector, TRIP, contains the bacteriophage T7 promoter for expression regulation of a therapeutic shRNA of interest. Furthermore, TRIP has the Inv locus from Yersinia pseudotuberculosis that encodes invasin, which permits natural noninvasive bacteria to enter &beta;1-integrin-positive mammalian cells and the HlyA gene from Listeria monocytogenes, which produces listeriolysin O. This enzyme allows the therapeutic shRNA to escape from entry vesicles within the cytoplasm of the target cell. TRIP constructs are introduced into a competent non-pathogenic Escherichia coli strain, which encodes T7 RNA polymerase necessary for the T7 promoter-driven synthesis of shRNAs. A well-characterized cancer-associated target molecule for different RNAi strategies is ABCB1 (MDR1/P-glycoprotein, MDR1/P-gp). This ABC-transporter acts as a drug extrusion pump and mediates the "classical" ABCB1-mediated multidrug resistance (MDR) phenotype of human cancer cells which is characterized by a specific cross resistance pattern. Different ABCB1-expressing MDR cancer cells were treated with anti-ABCB1 shRNA expression vector bearing E. coli. This procedure resulted in activation of the RNAi pathways within the cancer cells and a considerable down regulation of the ABCB1 encoding mRNA as well as the corresponding drug extrusion pump. Accordingly, drug accumulation was enhanced in the pristine drug-resistant cancer cells and the MDR phenotype was reversed. By means of this model the data provide the proof-of-concept that tkRNAi is suitable for modulation of cancer-associated factors, e.g. ABCB1, in human cancer cells.

For development of RNA interference (RNAi)-based therapies, a novel strategy was developed, transkingdom RNAi (tkRNAi). This technology uses non-pathogenic bacteria to produce and deliver therapeutic short hairpin RNA (shRNA) into target cells. Here, tkRNAi was successfully applied for reversal of classical ABCB1-mediated multidrug resistance (MDR) of cancer cells.

RNA interference (RNAi) represents a high effective mechanism for specific inhibition of mRNA expression. Besides its potential as a powerful laboratory ...

Direct Delivery of MIF Morpholinos Into the Zebrafish Otocyst by Injection and Electroporation Affects Inner Ear Development

In recent years, electroporation has become a popular technique for in vivo transfection of DNA, RNA, and morpholinos into various tissues, including the eye, brain, and somites of zebrafish. The advantage of electroporation over other methods of genetic manipulation is that specific tissues can be targeted, both spatially and temporally, for the introduction of macromolecules by the application of electrical current. Here we describe the use of electroporation for transfecting mif and mif-like morpholinos into the tissues of the developing inner ear of the zebrafish. In past studies, mif morpholino injected into embryos at the 1- to 8-cell stage resulted in widespread morphological changes in the nervous system and eye, as well as the ear. By targeting the tissues of the inner ear at later stages in development, we can determine the primary effects of MIF in the developing inner ear, as opposed to secondary effects that may result from the influence of other tissues. By using phalloidin and acetylated tubulin staining to study the morphology of neurons, neuronal processes, and hair cells associated with the posterior macula, we were able to assess the efficacy of electroporation as a method for targeted transfection in the zebrafish inner ear. The otic vesicles of 24hpf embryos were injected with morpholinos and electroporated and were then compared to embryos that had received no treatment or had been only injected or electroporated. Embryos that were injected and electroporated showed a decrease in hair cell numbers, decreased innervation by the statoacoustic ganglion (SAG) and fewer SAG neurons compared with control groups. Our results showed that direct delivery of morpholinos into otocysts at later stages avoids the non-specific nervous system and neural crest effects of morpholinos delivered at the 1-8 cell stage. It also allows examination of effects that are directed to the inner ear and not secondary effects on the ear from primary effects on the brain, neural crest or periotic mesenchyme.

A method to deliver morpholinos directly into the zebrafish otocyst at 24hpf has been developed. Using microinjection of morpholinos into the lumen of otic vesicle and electroporation to effect penetration, we were able to bypass the effect of morpholinos on the brain and obtain effects specific to the inner ear.

In recent years, electroporation has become a popular technique for in vivo transfection of DNA, RNA, and morpholinos into various tissues, including the ...

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 ...

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 ...

RNAi Screening to Identify Postembryonic Phenotypes in C. elegans

C. elegans has proven to be a valuable model system for the discovery and functional characterization of many genes and gene pathways1. More sophisticated tools and resources for studies in this system are facilitating continued discovery of genes with more subtle phenotypes or roles. 
Here we present a generalized protocol we adapted for identifying C. elegans genes with postembryonic phenotypes of interest using RNAi2. This procedure is easily modified to assay the phenotype of choice, whether by light or fluorescence optics on a dissecting or compound microscope. This screening protocol capitalizes on the physical assets of the organism and molecular tools the C. elegans research community has produced. As an example, we demonstrate the use of an integrated transgene that expresses a fluorescent product in an RNAi screen to identify genes required for the normal localization of this product in late stage larvae and adults. First, we used a commercially available genomic RNAi library with full-length cDNA inserts. This library facilitates the rapid identification of multiple candidates by RNAi reduction of the candidate gene product. Second, we generated an integrated transgene that expresses our fluorecently tagged protein of interest in an RNAi-sensitive background. Third, by exposing hatched animals to RNAi, this screen permits identification of gene products that have a vital embryonic role that would otherwise mask a post-embryonic role in regulating the protein of interest. Lastly, this screen uses a compound microscope equipped for single cell resolution.

RNAi Screening to Identify Postembryonic Phenotypes in C. elegans

We describe a sensitized method to identify postembryonic regulators of protein expression and localization in C. elegans using an RNAi-based genomic screen and an integrated transgene that expresses a functional, fluorescently tagged protein.

C. elegans has proven to be a valuable model system for the discovery and functional characterization of many genes and gene pathways1. More sophisticated ...