This work was supported in part by UCSD startup funds directed to O.S.A. and NSF/BIO grant 1645331 to J.L.R.

1. Nasonia rearing
<ol>
	<li>Set up ~20 mated females singularly in small glass test tubes plugged with cotton. 
	NOTE: To reduce rearing space, 10 x 85 cm, 4 mL tubes are optimal. Females are easily distinguished from males because of the larger wings and the presence of the ovipositor (Figure 1A). Detailed protocols for rearing N. vitripennis can also be found in other literature13,14,15.
	<ol>
		<li>Add two S. bullata hosts per tube, and maintain the wasps at 25 &#177; 1 &#176;C and 30% relative humidity, with a 12:12 light-dark cycle for 2-3 days. 
		&#8203;NOTE: Wasps can be maintained at lower temperature or room temperature; however, development will be slowed down.</li>
	</ol>
	</li>
	<li>After 2-3 days, with the help of a fine-tip paint brush, gently remove the females to avoid continuous oviposition and asynchrony in offspring development. 
	&#8203;NOTE: Removed females can optionally be re-hosted or stored at 5 &#176;C for up to a month.
	<ol>
		<li>Maintain the parasitized host at 25 &#176;C and 30% relative humidity, with a 12:12 light-dark cycle for 7 days if the desired injection stage requires white pupae; for 13 days if the required developmental stage is black pupae; or 14 days for young, newly emerged adults. 
		NOTE: Yellow and black pupae can also be stored at 5 &#176;C for up to one week. Longer storage will increase the frequency of diapause in larvae for the next generation, making the post-injection screening and establishment of mutant lines more difficult.</li>
		<li>Crack open the host puparium with a dissecting needle to recover N. vitripennis pupae and adults at the desired stage (Figure 1A). 
		NOTE: For injections at a particular adult age or for collecting virgin females, it is recommended to remove the dark pupae on day 13 from the host, separate by sex, and collect adults after emergence.</li>
	</ol>
	</li>
</ol>
2. Alignment of white and black pupae
<ol>
	<li>Prepare a glass slide by applying a line of school glue at the center. Spread the glue with a dissecting needle to obtain a thick layer (Figure 1B), which will be used to align the pupae. Let the glue dry for ~2 min before transferring the pupae. 
	NOTE: Do not overdry the glue, otherwise the pupae will not be properly attached to the slide and will slip over during the injections.</li>
	<li>Under a dissecting microscope, using the dissecting needle, apply glue on the back of the head of the pupa.</li>
	<li>Attach the pupa to the glue layer on the slide with its abdomen facing up. Align 20-30 pupae side by side on the slide (Figure 1C). 
	&#8203;NOTE: Avoid touching the abdomen with the glue, and avoid submerging the pupae into the glue; otherwise, the adults will not be able to emerge.</li>
	<li>Place the slide with the pupae in a Petri dish to let the glue dry for ~10 min. Before starting the injections, test the adherence of the pupae to the glue by pushing them gently with the dissecting needle. If most of the pupae are loosened, prepare a new slide. Alternatively, remove all the pupae that are loose, and proceed to inject those that are properly attached to the slide.</li>
</ol>
3. Needle preparation
<ol>
	<li>Load one capillary glass tube into a needle puller, and pull needles following the manufacturer&#39;s instructions. 
	NOTE: A P-1000 platinum filament needle puller and aluminum silicate glass capillaries are used in this demonstration. The operation manual for this instrument (see Table of Materials) explains how to properly load capillary glass tubes and set up programs on this instrument. Use the following parameters (Heat: 536; Pull: 80; Vel: 100; Delay: 70) on aluminum silicate glass capillaries to inject yellow pupae, black pupae, and adults. 
	In addition, the P-2000 needle puller was used to prepare quartz needles following the parameters (Heat: 805; Pull: 145; Vel: 50; Delay: 145). The manual provides instructions for loading capillary quartz tubes and setup of programs in this instrument. In the absence of a puller, customizable capillary glass is available.</li>
	<li>Load the needle with 5 &#181;L of the injection mixture (prepared in advance following previous protocols for RNAi10 or CRISPR/Cas9 ribonucleoprotein12 (RNP) injections in N. vitripennis) using microloader pipette tips. 
	NOTE: During the demonstration, the needle is loaded with double-stranded RNA (dsRNA) combined with red dye (see Table of Materials) for injection of white stage pupae and with BAPC-Cas9-sgRNA for injection of black pupae wasps and adults. These reagents will target the cinnabar gene. Wasps that are successfully knocked down/out will show red eyes instead of the wild type brown eyes.</li>
	<li>Open the needle, sliding the tip on a surface made from two overlapping slides (Figure 1D). Alternatively, open the tip of the needle with a pair of fine forceps to create a sharp edge or by slowly piercing the thorax or abdomen of the wasp.</li>
</ol>
4. Pupal injection with femtojet
<ol>
	<li>Place the slide with the aligned pupae under a dissecting microscope. Insert the needle into the femtojet&#39;s injection tube and tighten the grip head.</li>
	<li>Looking at the needle under the microscope, turn on the femtojet and set up Pc and Pi injection parameters by rotating the rotary knobs. 
	NOTE: A Pc value of 600 hPa is recommended for a continuous flow of the liquid. The Pc value depends on the opening of the needle. For needles prepared using the indicated parameters in the P-1000 needle puller, Pc values between 500 hPa and 700 hPa produce a constant flow. If lower values of Pc are required, this indicates that the opening may be too large and could damage the insects, whereas higher values indicate that the needle is still closed, or that the opening is too small.</li>
	<li>Carefully insert the needle between the 2 and 3 visible abdominal segments with a vertical angle of about 30&#176; (Figure 1E). Inject with a continuous flow until the whole abdomen turns pink in the case of white-stage pupae, or until the abdomen increases in size in the case of black pupae. Stop injecting when it is clear that the abdomen cannot take any more liquid, or when the liquid starts flowing out of the body. Move carefully to the next pupa and repeat these steps. 
	NOTE: Avoid touching and damaging the ovipositor with the needle during injection.</li>
	<li>Transfer the slide with the injected pupae to a Petri dish containing a paper towel soaked with deionized water to maintain humidity. Cover the dish with its lid, and place it at 25 &#176;C until wasp emergence (Figure 1F).</li>
</ol>
5. Pupal injection with aspirator tube
<ol>
	<li>Calculate the amount of injection mix using the factor: 4 pupae injected/1 &#181;L solution. 
	NOTE: A typical injection of 20-30 pupae requires ~ 5-8 &#181;L of ribonucleoprotein (RNP) complex-saponin mix or RNP with BAPC12.</li>
	<li>Align the pupae, as suggested in section 2, and place one slide with the aligned pupae under a dissecting microscope (Figure 1E).</li>
	<li>Take one of the capillary needles and break the tip between two glass slides, as indicated in step 3.4 (Figure 1D). Ensure that the needle tip is open enough to allow the injection solution to go out by blowing air with the mouth, but not too open to avoid losing liquid and damaging the pupae (Figure 1E). 
	NOTE: This step is critical because the user will be using air from the mouth to push the injection mixture into the insect hemolymph. It is better to practice so as to ascertain what type of needle aperture is optimal for the viscosity of a particular solution mix. Five sets of injections of 20 pupae per set may be enough to get used to the mouth injection system.</li>
	<li>Load one needle with the ribonucleoprotein solution12 using a microloading tip, and insert the needle into the connector pack of the aspirator tube assembly. 
	NOTE: During the demonstration, the needle is loaded with BAPC-Cas9-sgRNA targeting the Nasonia cinnabar gene12.</li>
	<li>Carefully insert the needle between the 2 and 3 visible abdominal segments with a vertical angle of about 30&#176; (Figure 1E). Inject with a continuous flow until the whole abdomen turns pink in the case of white-stage pupae or until the abdomen increases in size in the case of black pupae. Stop injecting when it is clear that the abdomen cannot take any more liquid or when the liquid starts flowing out of the body. Move carefully to the next pupa and repeat these steps 
	NOTE: Avoid touching and damaging the ovipositor with the needle during injection.</li>
	<li>Transfer the slide with the injected pupae to a Petri dish containing a paper towel soaked with deionized water to maintain humidity. Cover the dish with its lid and place it at 25 &#176;C until wasp emergence (Figure 1F).</li>
</ol>
6. Adult injection with aspirator tube
<ol>
	<li>For adult preparation, separate groups of 20 virgin females in a clean small test tube with a fine paintbrush. Place the tube on ice for 5 min until the females are anesthetized. Alternatively, females can be anesthetized and injected using CO2. 
	NOTE: Ice is preferable as an anesthetic as adult wasps are cold-tolerant and recover easily after the injections. Females can take up more injected liquid than the pupae: 1 &#181;L of solution can be used for three females instead of for four pupae. Prepare volumes accordingly.</li>
	<li>Prepare an ice bucket, placing one glass slide on top of the ice. Align the females side by side on the cold slide with abdomens facing up (Figure 1E) under a dissecting scope.</li>
	<li>Load one needle with the ribonucleoprotein solution12 using a microloading tip, and insert the needle into the connector pack of the aspirator tube assembly. Support the females from the opposite side with blunted dissecting needles while injecting slowly into the abdomen from the other side. Avoid touching the ovipositor, this will severely damage this critical reproductive structure (Figure 1E). 
	NOTE: Either orientation is acceptable, depending on operator preference.</li>
	<li>Carefully insert the needle between the 2 and 3 visible abdominal segments with a vertical angle of about 30&#176; (Figure 1E). Inject the solution into the female abdomen, stopping when no more liquid can enter, or when leaking of surplus solution is observed. Move carefully to the next wasp&#160;and repeat these steps 
	NOTE: Inject slowly, leaving the needle inside the abdomen for about 3 s before removing it very slowly. This will help to adjust the internal liquid pressure and avoid solution leaking from the injection site following needle removal.</li>
	<li>When finished, gently transfer single injected females to a new tube with one host using a paintbrush. Leave to recover at room temperature for approximately 1 h, confirming survival, and then return the tubes to the rearing incubator. 
	&#8203;NOTE: During the demonstration, the needle is loaded with BAPC-Cas9-sgRNA targeting the Nasonia cinnabar gene12.</li>
</ol>
7. Post-injection care and mutant screening
<ol>
	<li>After adult eclosion from the injected pupae, place single wasps into a glass tube plugged with cotton and insert one S. bullata host (Figure 1G). In contrast, place injected females in individual tubes with hosts immediately after injection.</li>
	<li>For CRISPR/Cas9 knockout experiments, allow females to parasitize the hosts for one day, and replace the host each day for three consecutive days. 
	NOTE: Owing to the haplodiploid sex determination system in N. vitripennis16, virgin females will produce haploid male broods, which facilitates the detection of mutations. For knockdown via RNAi of male-specific genes, the injected individuals can be singularly mated with wild type individuals and allowed to parasitize hosts. Use one host per day, and replace the hosts based on the experimental setup.</li>
	<li>Place the parasitized hosts at 25 &#176;C until emergence of the G0 male offspring (for ~13-14 days).</li>
	<li>Under a dissecting microscope, screen G0 males for the mutated phenotypes. The cinnabar gene is responsible for eye pigmentation12: wasps with brown eyes are wild type, and wasps with bright red eyes or variations between red and brown eyes are mutants (Figure 1H).</li>
</ol>
8. Post-injection crosses and rearing
<ol>
	<li>Place all the mutant G0 males with red eyes (the phenotypically disrupted cinnabar12 gene) with wild type virgin females for 1-2 days (Figure 2A). 
	NOTE: If the disrupted gene does not confer a visible phenotype, polymerase chain reaction (PCR) followed by sequencing of the target gene is required to identify the mutant animals. Before obtaining the DNA from G0 males for PCR, it would be ideal to mate them with wild type virgin females. Alternatively, extract DNA from a leg of a G0 male to identify mutants, and mate only those with verified mutations.</li>
	<li>Add two S. bullata hosts per female, and allow to oviposit at 25 &#176;C and 30% relative humidity with a 12:12 light-dark cycle. Replace the hosts every 2-3 days.
	<ol>
		<li>Store parasitized hosts at 25 &#176;C and 30% relative humidity, with a 12:12 light-dark cycle for ~10 days.</li>
		<li>After ~10 days, crack open the parasitized host with a dissecting needle, and remove each N. vitripennis pupa from the host.</li>
		<li>With the help of a fine-tip paintbrush, select female pupae (Figure 1A). 
		&#8203;NOTE: Females are diploid and heterozygous, and thus the eye color will be wild type.</li>
	</ol>
	</li>
	<li>Place 15-20 female pupae in glass tubes at 25 &#176;C until emergence. After emergence, add ~20 S. bullata hosts and let the virgin G1 females parasitize the hosts for 2-3 days. 
	NOTE: These females will produce 50%&#160;G2 mutant males.
	<ol>
		<li>Store parasitized hosts at 25 &#176;C and 30% relative humidity, with a 12:12 light-dark cycle until G2 male adult emergence.</li>
		<li>Place the remaining G1 pupae at 5 &#176;C for 8-10 days . After this period, remove them and place them at 25 &#176;C and 30% relative humidity, with a 12:12 light-dark cycle until emergence (Figure 2A).</li>
	</ol>
	</li>
	<li>After emergence of the G2 males in step 8.3.1, screen for the presence of red-eyed mutant phenotype. With the help of a fine-tip paintbrush, separate red-eyed males from wild type males. 
	&#8203;NOTE: The presence of red-eyed wasps in this generation (G2, Figure 2A) indicates that gene editing occurred in the germline, and that the mutation is hereditary.</li>
	<li>Place red-eyed males with the G1 females newly emerged in step 8.3.2 (Figure 2A), and let them mate for 1-2 days. Add 2 S. bullata hosts per female, and place them at 25 &#176;C and 30% relative humidity with a 12:12 light-dark cycle for 10-11 days.
	<ol>
		<li>After 10-11 days, crack open the parasitized host with a dissecting needle, and remove each N. vitripennis pupa from the host (Figure 2A).</li>
		<li>With the help of a fine-tip paintbrush, separate red-eyed wasps (males and females) from wild type (Figure 1H). Continue rearing of red-eyed wasps together in glass tubes, and do not mix with the wild type wasps. 
		NOTE: At this stage (G3, Figure 2A), haploid males and diploid females carry and show phenotypes for the cinnabar mutations. If the affected gene encodes for an invisible phenotype, cross G0 male singularly with one wild type female for ~1 day. Then, remove the male for molecular characterization of the mutation by PCR and sequencing, and let the female parasitize a host. Continue the crossing scheme only with the offspring of a confirmed mutant male (Figure 2B). If the goal is the knockdown by RNAi of a candidate gene, perform the desired functional assay (such as lethality, sterility, or diapause induction assay3,7) directly using individuals injected with dsRNA. As RNAi is transient, it is not possible to generate a colony with the desired phenotype (Figure 2C)3,7.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61892/61892fig1v2.jpg" /> 
Figure 1: Timeline for pupal and adult&#160;Nasonia vitripennis&#160;microinjection.&#160;(A) Male and female white (top) and black (bottom) pupae are collected, (B-C) aligned and glued to a glass slide for injection. (D) Capillary needle is prepared and opened, sliding it on two overlapping slides. (E) Needle is attached either to the Femtojet (top) or to an aspirator tube (bottom) for injections.(F) The injected pupae on the slide are transferred to a Petri dish with a wet tissue on the bottom to keep humidity. Upon emergence, (G) females are placed singularly in small glass tubes and allowed to oviposit on&#160;Sarcophaga&#160;pupa. (H) Screening of the offspring to detect mutants.&#160;<a href="https://www.jove.com/files/ftp_upload/61892/61892fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61892/61892fig2v2.jpg" /> 
Figure 2: Crossing scheme after injection. Schematic representation of crossing scheme procedure in the case of CRISPR/Cas9-mediated gene editing of genes&#160;(A)&#160;that induce a visible phenotype and&#160;(B)&#160;that do not confer visible phenotype.&#160;(C)&#160;Schematic representation of RNAi screening procedure. Abbreviations:&#160;cin&#160;=&#160;cinnabar; PCR = polymerase chain reaction.&#160;<a href="https://www.jove.com/files/ftp_upload/61892/61892fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNAi+analysis+of+Deformed,+proboscipedia+and+Sex+combs+reduced+in+the+milkweed+bug+Oncopeltus+fasciatus:+novel+roles+for+Hox+genes+in+the+hemipteran+head.">RNAi analysis of Deformed, proboscipedia and Sex combs reduced in the milkweed bug Oncopeltus fasciatus: novel roles for Hox genes in the hemipteran head.</a> Development. 127 (17), 3683-3694 (2000).</li><li>Dzitoyeva, S., Dimitrijevic, N., Manev, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intra-abdominal+injection+of+double-stranded+RNA+into+anesthetized+adult+Drosophila+triggers+RNA+interference+in+the+central+nervous+system.">Intra-abdominal injection of double-stranded RNA into anesthetized adult Drosophila triggers RNA interference in the central nervous system.</a> Molecular Psychiatry. 6 (6), 665-670 (2001).</li><li>Chaverra-Rodriguez, D., 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=Targeted+delivery+of+CRISPR-Cas9+ribonucleoprotein+into+arthropod+ovaries+for+heritable+germline+gene+editing.">Targeted delivery of CRISPR-Cas9 ribonucleoprotein into arthropod ovaries for heritable germline gene editing.</a> Nature Communications. 9 (1), 3008 (2018).</li><li>Macias, V. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cas9-mediated+gene-editing+in+the+malaria+mosquito+anopheles+stephensi+by+ReMOT+control.">Cas9-mediated gene-editing in the malaria mosquito anopheles stephensi by ReMOT control.</a> G3. 10 (4), 1353-1360 (2020).</li><li>Heu, C. C., McCullough, F. M., Luan, J., Rasgon, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9-based+genome+editing+in+the+silverleaf+whitefly+(Bemisia+tabaci).">CRISPR/Cas9-based genome editing in the silverleaf whitefly (Bemisia tabaci).</a> CRISPR Journal. 3 (2), 89-96 (2020).</li><li>Shirai, Y., Daimon, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutations+in+cardinal+are+responsible+for+the+red-1+and+peach+eye+color+mutants+of+the+red+flour+beetle+Tribolium+castaneum.">Mutations in cardinal are responsible for the red-1 and peach eye color mutants of the red flour beetle Tribolium castaneum.</a> Biochemical and Biophysical Research Communications. 529 (2), 372-378 (2020).</li><li>Hunter, W. B., Gonzalez, M. T., Tomich, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BAPC-assisted+CRISPR/Cas9+system:+targeted+delivery+into+adult+ovaries+for+heritable+germline+gene+editing+(Arthropoda:+Hemiptera).">BAPC-assisted CRISPR/Cas9 system: targeted delivery into adult ovaries for heritable germline gene editing (Arthropoda: Hemiptera).</a> bioRxiv. , 478743 (2018).</li><li>Chaverra Rodriguez, D., Bui, M., Li, M., Raban, R., Akbari, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developing+genetic+tools+to+control+ACP.">Developing genetic tools to control ACP.</a> Citrus Research Board Citrograph Magazine. 11 (1), 64-68 (2020).</li></ol>

JLR and DCR have filed for provisional patent protection on ReMOT Control technology. O.S.A is a founder of Agragene, Inc., has an equity interest, and serves on the company&#39;s Scientific Advisory Board. All other authors declare no competing interests.

With the recent increased use of Nasonia vitripennis as a model organism for various biological questions2,3,7,17, there is a need to develop and optimize injection methods to enable a simplified and efficient protocol for the functional analysis of N. vitripennis genes. The current methods involving embryonic microinjection of gene editing reagents are challenging<sup class="...

CRISPR/Cas9 gene editing is a powerful technology for functional genetic studies, especially in many rising model organisms such as the jewel wasp, Nasonia vitripennis. The ease of rearing and the availability of a complete genome makes the jewel wasp an important experimental system for elucidating the molecular mechanisms of different biological processes. For example, N. vitripennis has recently been used to unravel the epigenetic basis of the haplodiploid sex determination system1,2, the biology of B-chromosomes3,4,5,6, and the genetic basis for circadian and seasonal regulation7,8. Some of the features that make N. vitripennis amenable to work with include short generation time (~2 weeks at 25 &#176;C), high reproduction rates, easy sex separation at the pupal stage, and the ability to diapause and store strains at 4 &#176;C. The life cycle begins with female wasps parasitizing the pupae of the blowfly, Sarcophaga bullata. Through their ovipositor, females lay up to 50 eggs in the pupal case of the blowfly. Eggs develop into larvae that feed on the S. bullata pupa, continue to develop over the next several days, and then pupate, followed by adult eclosion and emergence from the host puparium9.
Molecular tools to perform functional genetic studies in N. vitripennis, such as RNA interference (RNAi)10 and CRISPR/Cas911,12, are available, but are limited, primarily due to difficulties in performing embryonic microinjections13. As N. vitripennis eggs require a pupal host for development, egg manipulation is very challenging. Pre-blastoderm stage embryos must be collected from the host blowfly pupae, quickly microinjected, and immediately transferred back to the host for development13. These steps require precision and specialized training to avoid damaging the microinjected embryos or the pupal hosts13. Moreover, the eggs are very small and fragile, especially after microinjections, with a very viscous cytoplasm causing a continuous clogging of the injecting needle13. These features make embryonic microinjections exceptionally challenging, requiring highly trained operators and specialized equipment that is absent in most N. vitripennis laboratories.
The optimization of alternative injection methods for the delivery of CRISPR reagents would contribute to the consolidation of N. vitripennis as a model organism. The manipulation of pupae and adults is less challenging than manipulating embryos and can be accomplished with a basic injection setup. Here, two protocols are described for injection of pupae and adults: one involving specialized equipment for injections, and the other involving the use of an aspirator tube assembly fitted with a glass capillary needle. The use of an aspirator tube is particularly suited for laboratories that do not have access to specialized equipment for embryo microinjections. Efficient injections of different developmental stages of N. vitripennis, including white or black pupae and adult wasps, are demonstrated. Wasps at the white pupal stage are particularly suited for RNAi-mediated knockdown experiments. Although RNAi in Nasonia was first described by Lynch and Desplan in 200610, there is no visual procedure available for how RNAi injections are performed. RNAi was recently used to discover the haploidizer gene of the B-chromosome PSR (Paternal Sex Ratio)3 and to study the involvement of the clock gene, period, in N. vitripennis biological rhythms7.
Black pupae and adult wasps can be used to induce CRISPR/Cas9 germline gene editing using ReMOT Control (Receptor-Mediated Ovary Transduction of Cargo) and BAPC (Branched Amphiphilic Peptide Capsules) protocols. These two ovary delivery methods have been recently described to be effective in Nasonia for generating germline mutations in the target gene, cinnabar7,12. Here, a simplified protocol is provided for injections including a visual procedure of a step-by-step methodology for both pupal and adult injections that can be utilized to generate functional genetics studies in Nasonia and likely in other parasitoid wasps, without requiring specialized equipment and bypassing embryonic microinjections.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100 x 15 mm Stackable Petri Dishes, Polystyrene, Mono, Sterile</td>
			<td>Sigma</td>
			<td>960-97693-083</td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminosilicate glass capillary tubing 1 mm(outside diameter) x 0.58 mm (inner diameter)</td>
			<td>Sutter Instruments</td>
			<td>BF100-58-10</td>
			<td>Can also use Borosilicate or Quartz</td>
		</tr>
		<tr>
			<td>Aspirator tube assemblies for calibrated microcapillary pipettes</td>
			<td>Sigma</td>
			<td>A5177-5EA</td>
			<td></td>
		</tr>
		<tr>
			<td>BAPC</td>
			<td>phoreus biotech</td>
			<td>BAPtofect-25 0.5 mg Kit</td>
			<td></td>
		</tr>
		<tr>
			<td>Cas9 Protein with NLS</td>
			<td>PNABio</td>
			<td>CP01</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting needle</td>
			<td>VWR</td>
			<td>10806-330</td>
			<td></td>
		</tr>
		<tr>
			<td>DNase/RNase-Free distilled Water</td>
			<td>Invitrogen</td>
			<td>10977-015</td>
			<td></td>
		</tr>
		<tr>
			<td>Femtojet Express programmable microinjector</td>
			<td>Eppendorf</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Femtotips Microloader tips</td>
			<td>Fisher Scientific</td>
			<td>E5242956003</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine-tip paintbrush</td>
			<td>ZEM</td>
			<td>2595</td>
			<td></td>
		</tr>
		<tr>
			<td>Flesh fly pupae,&#160;Sarcophaga bullata</td>
			<td>Ward&#39;s Science</td>
			<td>470180-392</td>
			<td></td>
		</tr>
		<tr>
			<td>Food colorant dye</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Test Tubes</td>
			<td>Fisher Scientific</td>
			<td>982010</td>
			<td></td>
		</tr>
		<tr>
			<td>Glue</td>
			<td>Elmer&#39;s</td>
			<td></td>
			<td>washable, no toxic school glue</td>
		</tr>
		<tr>
			<td>Micropipette Puller</td>
			<td>Sutter Instruments</td>
			<td>P-1000 or P-2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Slides</td>
			<td>Fisherbrand</td>
			<td>12-550-A3</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereo Microscope</td>
			<td>Olympus</td>
			<td>SZ51</td>
			<td></td>
		</tr>
	</tbody>
</table>

pupal and adult injections for rnai and crispr gene editing in nasonia vitripennis

The jewel wasp, Nasonia vitripennis, has become an efficient model system to study epigenetics of haplo-diploid sex determination, B-chromosome biology, host-symbiont interactions, speciation, and venom synthesis. Despite the availability of several molecular tools, including CRISPR/Cas9, functional genetic studies are still limited in this organism. The major limitation of applying CRISPR/Cas9 technology in N. vitripennis stems from the challenges of embryonic microinjections. Injections of embryos are particularly difficult in this organism and in general in many parasitoid wasps, due to small embryo size and the requirement of a host pupa for embryonic development. To address these challenges, Cas9 ribonucleoprotein complex delivery into female&#160;ovaries by adult injection, rather than embryonic microinjection, was optimized, resulting in both somatic and heritable germline edits. The injection procedures were optimized in pupae and female wasps using either ReMOT Control (Receptor-Mediated Ovary Transduction of Cargo) or BAPC (Branched Amphiphilic Peptide Capsules). These methods are shown to be effective alternatives to embryo injection, enabling site-specific and heritable germline mutations.

This paper presents two easy methods for pupal and adult microinjection, either using a femtojet or an aspirator tube. The first method allows a more precise injection of liquid, which is important for RNAi consistency, whereas the second one allows the injection of larger amounts of liquid into Nasonia pupae or adults.
Representative results presented in Table 1 show good survival rates (from 20% to 89...

Watch this Scientific Journal Video about Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis at JoVE.com

Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis

Here, we describe methods for efficient pupal and adult injections in Nasonia vitripennis as accessible alternatives to embryo microinjection, enabling functional analysis of genes of interest using either RNA-silencing via RNA interference (RNAi) or gene knockout via CRISPR/Cas9 genome editing.

Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis

The jewel wasp, Nasonia vitripennis, has become an efficient model system to study epigenetics of haplo-diploid sex determination, B-chromosome biology, ...

pupal-adult-injections-for-rnai-crispr-gene-editing-nasonia

Research

JoVE Journal

Biology

5.1K Views.  University of California, San Diego. Here, we describe methods for efficient pupal and adult injections in Nasonia vitripennis as accessible alternatives to embryo microinjection, enabling functional analysis of genes of interest using either RNA-silencing via RNA interference (RNAi) or gene knockout via CRISPR/Cas9 genome editing. 

Video: Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis

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 ...

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of muscle specification, formation and function in model systems has provided valuable insight into muscle physiology. Therefore, identifying and characterizing molecular mechanisms that underlie muscle damage is critical. The structure of adult Drosophila multi-fiber muscles resemble vertebrate striated muscles 1 and the genetic tractability of Drosophila has made it a great system to analyze dystrophic muscle morphology and characterize the processes affecting muscular function in ageing adult flies 2. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. These preparations allow for the tissue to be stained with classical histological stains and labeled with protein detecting dyes, and specifically cryosections are ideal for immunohistochemical detection of proteins in intact muscles. This allows for analysis of muscle tissue structure, identification of morphological defects, and detection of the expression pattern for muscle/neuron-specific proteins in Drosophila adult muscles. These techniques can also be slightly modified for sectioning of other body parts.

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Identification of mechanisms underlying muscle damage is crucial. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. This allows analysis of muscle morphology and localization of protein and other muscle cell components.

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of ...

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-mediated Gene Knockdown and In Vivo Diuresis Assay in Adult Female Aedes aegypti Mosquitoes

This video protocol demonstrates an effective technique to knockdown a particular gene in an insect and conduct a novel bioassay to measure excretion rate. This method can be used to obtain a better understanding of the process of diuresis in insects and is especially useful in the study of diuresis in blood-feeding arthropods that are able to take up huge amounts of liquid in a single blood meal. 
This RNAi-mediated gene knockdown combined with an in vivo diuresis assay was developed by the Hansen lab to study the effects of RNAi-mediated knockdown of aquaporin genes on Aedes aegypti mosquito diuresis1. 
The protocol is setup in two parts: the first demonstration illustrates how to construct a simple mosquito injection device and how to prepare and inject dsRNA into the thorax of mosquitoes for RNAi-mediated gene knockdown. The second demonstration illustrates how to determine excretion rates in mosquitoes using an in vivo bioassay.

RNAi-mediated Gene Knockdown and In Vivo Diuresis Assay in Adult Female Aedes aegypti Mosquitoes

In this protocol we combine RNAi-mediated gene silencing with an in vivo diuresis assay to study the effects knockdown of genes of interest has on mosquito fluid excretion.

This video protocol demonstrates an effective technique to knockdown a particular gene in an insect and conduct a novel bioassay to measure excretion ...

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter implantation. The technical limitation of this surgical technique lies in the finesse of the hands. The injection is rapid, especially for a trained experimenter, and since tissue disruption with this technique is minimal, repeated injections are possible; moreover immune reaction to foreign tools (e.g. catheter) does not occur, thereby giving a better and more specific read out of spinal cord modulation. Since the application of the substance is largely limited to the target region of the spinal cord, drugs do not need to be applied in large dosages, and more importantly unwanted effects on other tissue, as observed with a systemic delivery, could be circumvented1,2. Moreover, we combine this technique with in vivo transfection of nucleic acid with the help of polyethylenimine (PEI) reagent3, which provides tremendous versatility for studying spinal functions via delivery of pharmacological agents as well as gene, RNA, and protein modulators.

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a simple and rapid technique of intrathecal needle puncture for a localized transfection of siRNA in the lumbar spinal cord in mouse under short lasting light anesthesia.

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter ...

Intravenous Injections in Neonatal Mice

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are technically feasible and routine. However, some mouse models can have early onset of disease with a rapid progression that makes administration of potential therapies difficult. The temporal (or facial) vein is just anterior to the ear bud in mice and is clearly visible for the first two days after birth on either side of the head using a dissecting microscope. During this window, the temporal vein can be injected with volumes up to 50 &#956;l. The injection is safe and well tolerated by both the pups and the dams. A typical injection procedure is completed within 1-2 min, after which the pup is returned to the home cage. By the third postnatal day the vein is difficult to visualize and the injection procedure becomes technically unreliable. This technique has been used for delivery of adeno-associated virus (AAV) vectors, which in turn can provide almost body-wide, stable transgene expression for the life of the animal depending on the viral serotype chosen.

Animal models of pediatric disease can experience early onset and aggressive disease progression. Clinically relevant therapy delivery to young mouse models can be technically difficult. This protocol describes a non-invasive intravenous injection method for newborn mice within the first two postnatal days of life.

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are ...

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these independently derived phenotypes, which share a common genetic architecture. This is perhaps best exemplified by the numerous convergent features of gymnotiforms and mormyrids, two species-rich teleost clades that produce and detect weak electric fields and are called weakly electric fish. In the 50 years since the discovery that weakly electric fish use electricity to sense their surroundings and communicate, a growing community of scientists has gained tremendous insights into evolution of development, systems and circuits neuroscience, cellular physiology, ecology, evolutionary biology, and behavior. More recently, there has been a proliferation of genomic resources for electric fish. Use of these resources has already facilitated important insights with regards to the connection between genotype and phenotype in these species. A major obstacle to integrating genomics data with phenotypic data of weakly electric fish is a present lack of functional genomics tools. We report here a full protocol for performing CRISPR/Cas9 mutagenesis that utilizes endogenous DNA repair mechanisms in weakly electric fish. We demonstrate that this protocol is equally effective in both the mormyrid species Brienomyrus brachyistius and the gymnotiform Brachyhypopomus gauderio by using CRISPR/Cas9 to target indels and point mutations in the first exon of the sodium channel gene scn4aa. Using this protocol, embryos from both species were obtained and genotyped to confirm that the predicted mutations in the first exon of the sodium channel scn4aa were present. The knock-out success phenotype was confirmed with recordings showing reduced electric organ discharge amplitudes when compared to uninjected size-matched controls.

Here, a protocol is presented to produce and rear CRISPR/Cas9 genome knockout electric fish. Outlined in detail are the required molecular biology, breeding, and husbandry requirements for both a gymnotiform and a mormyrid, and injection techniques to produce Cas9-induced indel F0 larvae.

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these ...

Nuclei Isolation from Adult Mouse Kidney for Single-Nucleus RNA-Sequencing

The kidneys regulate diverse biological processes such as water, electrolyte, and acid-base homeostasis. Physiological functions of the kidney are executed by multiple cell types arranged in a complex architecture across the corticomedullary axis of the organ. Recent advances in single-cell transcriptomics have accelerated the understanding of cell type-specific gene expression in renal physiology and disease. However, enzyme-based tissue dissociation protocols, which are frequently utilized for single-cell RNA-sequencing (scRNA-seq), require mostly fresh (non-archived) tissue, introduce transcriptional stress responses, and favor the selection of abundant cell types of the kidney cortex resulting in an underrepresentation of cells of the medulla.
Here, we present a protocol that avoids these problems. The protocol is based on nuclei isolation at 4 &#176;C from frozen kidney tissue. Nuclei are isolated from a central piece of the mouse kidney comprised of the cortex, outer medulla, and inner medulla. This reduces the overrepresentation of cortical cells typical for whole-kidney samples for the benefit of medullary cells such that data will represent the entire corticomedullary axis at sufficient abundance. The protocol is simple, rapid, and adaptable and provides a step towards the standardization of single-nuclei transcriptomics in kidney research.

Here, we present a protocol to isolate high-quality nuclei from frozen mouse kidneys that improve the representation of medullary kidney cell types and avoids the gene expression artifacts from enzymatic tissue dissociation.

The kidneys regulate diverse biological processes such as water, electrolyte, and acid-base homeostasis. Physiological functions of the kidney are ...

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

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

Effective Oral RNA Interference RNAi Administration to Adult Anopheles gambiae Mosquitoes

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