This work was supported by a generous University of California, Riverside (UCR) laboratory start-up fund to O.S.A, a USDA National Institute of Food and Agriculture (NIFA) Hatch project (1009509) to O.S.A.

1. N. Vitripennis Colony Rearing
<ol>
	<li>Set up several colonies (~ 3-5) by placing 200-500 N. vitripennis adults (with a 3:1 ratio of female:male) in bug dorm cages. 
	NOTE: Alternative to using bug dorms, wasps can also be propagated in multiple, small glass test tubes plugged with cotton with approximately 15-20 wasps/vial.
	<ol>
		<li>Maintain the wasps at 25 &#177; 1 &#176;C with 30% relative humidity and a 12:12 light dark cycle, feeding them daily with small droplets of 1:10 (v/v) sucrose/water solution. 
		NOTE: Propagation of wasp colonies and collections can also be conducted at room temperature without controlled humidity; however, temperatures lower than 25 &#176;C will slightly lower the developmental timing of the embryos.</li>
	</ol>
	</li>
	<li>Allow wasps to mate for at least 4 days prior to injections.
	<ol>
		<li>For the first two days, allow mated females to feed on fresh S. bullata pupae, as well as small droplets of a 1:10 (v/v) sucrose/water solution.</li>
		<li>For the following two days, remove the blowfly pupae to deprive the female wasps of an oviposition site, making them very gravid.</li>
	</ol>
	</li>
</ol>
2. Collection and Alignment of N. Vitripennis Pre-blastoderm Stage Embryos
<ol>
	<li>Allow female wasps to parasitize (oviposit embryos) into young hosts (S. bullata pupae). To keep hosts fresh, store hosts in 4 &#176;C immediately after obtaining them and only remove when needed. 
	NOTE: Young hosts can be determined by having a red colored puparium whereas older hosts have a darker colored puparium (Figure 1A).
	<ol>
		<li>Place individual fresh S. bullata pupae into a foam stopper with a pupae-sized hole cut into the center. Expose only ~ 0.2 cm of the anterior end (preferred oviposition site) of the host for maximum concentration of parasitization and easier embryo collection. 
		NOTE: The anterior end is rounded, while the posterior end is thicker and contains a &#39;crater-like&#39; opening (Figure 1B).</li>
		<li>Place host pupae (roughly 5 host pupae per 100 wasps) in this arrangement into the cage and wait roughly 45 min to allow wasps to parasitize them (Figure 2- ii). 
		NOTE: It is very important that the embryos are as young as possible, ideally within the first hour of being oviposited, to ensure that they are in the pre-blastoderm stage. Old embryos (&#62; 1.5 h) should not be injected.</li>
		<li>Remove parasitized hosts by retrieving them by hand and gently brushing/blowing off any residing wasps. Replace them with fresh hosts after every ~ 15 min to continue collecting early staged eggs during injections.</li>
	</ol>
	</li>
	<li>Remove the freshly parasitized hosts from cage by hand. Under a dissecting microscope, using forceps, carefully peel away the posterior end (0.4 cm) of the exterior puparium (pupal shell) to expose the N. vitripennis eggs (Figure 2- iii). 
	NOTE: Care must be taken to avoid puncturing the pupal skin; if this happens, the hemolymph will moisten the wasp embryos, which will be very difficult to remove for microinjection.</li>
	<li>Use liquid adhesive to glue a coverslip onto a clean glass slide to prepare an embryo alignment slide (Figure 3). 
	NOTE: A high performance instant adhesive is used to increase bond strength and drying speed, however any glue that can keep the coverslip from moving around on the slide is acceptable.</li>
	<li>Brush-off the embryos from the host carefully with a wet fine-tip paintbrush, making sure not to damage the soft pupal skin of the host. 
	NOTE: An &#39;embryo pick,&#39; which is essentially a half of a pair of ultrafine forceps, can be used instead of a paintbrush.</li>
	<li>Transfer ~ 20 embryos one-by-one onto the slide, immediately adjacent to the side of the coverslip with a wet paintbrush. Orient the anterior end of the embryo against the cover slide&#39;s edge so that the posterior end of the embryos is pointed in the same direction (Figure 2- iv). This will allow for higher precision of injecting into the same position among all embryos. 
	NOTE: This improved technique does not require double-sided sticky tape to fix embryos to the slide as described previously21,22. Additionally, do not desiccate embryos, or cover the eggs with halocarbon oil during injection as described previously for N. vitripennis embryo microinjection22.</li>
</ol>
3. Needle Preparation for Microinjections
<ol>
	<li>Place an aluminosilicate capillary glass into a needle puller. 
	NOTE: A good needle is critical for successful N. vitripennis embryos injections. The injection needles should have a sharp and fine-tip to allow easy penetration through the chorion. Quartz and borosilicate capillary needles also can be used (Figure 4).</li>
	<li>Set the Heat to 605, the Velocity to 130, the Delay to 80, the Pull to 70, and the Pressure to 500 on a needle puller. 
	NOTE: Additional needle puller settings for pulling quartz and Borosilicate needles can be seen in Table 1; the parameters may vary for different pullers and filaments.</li>
	<li>Activate the needle puller to create the correct needle. Repeat as needed for production of multiple needles. 
	NOTE: Pulled needles can be stored by placing them into a Petri dish containing several parallel rolls of commercial modeler&#39;s clay.</li>
	<li>Bevel the needle tip by slightly touching the tip to the diamond abrasive plate around 10 s at 25 &#176;C (Figure 2- iii). 
	NOTE: The injection needles benefit from beveling by promoting entry into the embryos with minimal damage while simultaneously enhancing the flow of regents through the needle.</li>
</ol>
4. Embryo Microinjection
<ol>
	<li>Prepare the injection mixture consisting of genome modification reagents (e.g., transposon + helper transposase, or CRIPSR reagents, etc.), and keep it on ice.</li>
	<li>Load the injection needle with 2 &#181;L of injection mixture by using microloader tips. 
	NOTE: The injection mixture demonstrated in the protocol consists of single-guide RNA (sgRNA)(s) and recombinant Cas9 protein mixed in H2O.</li>
	<li>Place the glass slide with the lined embryos onto the stage of a compound microscope.</li>
	<li>Carefully insert the needle into the posterior end of the embryo with a vertical angle of 25-35 &#176;. Inject 1-5 pL of injection mixture (around 2-10% of the embryo&#39;s volume) and ensure that the embryo appears to swell slightly as the mixture is injected into it (Figure 2- iv). 
	NOTE: If too much injection mixture is injected into the embryo it may burst dispelling the cytoplasm from the embryo. Additionally, improperly beveled or broken needles with too large of an opening may also cause embryo rupture.
	<ol>
		<li>Try re-beveling needles if clogging occurs. The cytoplasm of N. vitripennis embryos is unusually viscous and sticky, which leads to frequent needle clogging (1/25 injections). 
		NOTE: It is important to keep the embryos moist during the injection period by regularly adding de-ionized water using the paintbrush. The amount of water on the brush is key to move embryos around and align with ease. Too much water results in embryos floating and too little water makes them difficult to move around.</li>
		<li>Inject ~ 20-40 eggs at a time (should take roughly 10 min), then stop. Transfer the injected eggs with a wet paintbrush by lightly touching the injected eggs and place them into a host. Then continue injecting again using a fresh newly laid batch of eggs (&#62; 1 h old). 
		NOTE: Ideally this protocol is most effective if one person is continuously collecting and lining up eggs, while another person is injecting the genome modification components and transplanting injected embryos to host pupae.</li>
	</ol>
	</li>
</ol>
5. Transplanting Injected G0 N. vitripennis Embryos onto Pre-stung Host
<ol>
	<li>Carefully place the injected G0 embryos onto a previously stung S. bullata pupae (Figure 2- vi) with a wet paintbrush after microinjection. 
	NOTE: The host pupae used to collect embryos for microinjection will work for this purpose. Also, despite significant effort, there is currently no available artificial diet that can be used22.
	<ol>
		<li>Place up to 40 injected embryos one at a time onto a single pre-stung host to avoid overcrowding and larval competition. 
		NOTE: Injection will cause damage to the embryos; careless transplantation of injected embryos to the host pupae may squeeze the injected component or the yolk out, and lead to the death of the embryos.</li>
	</ol>
	</li>
	<li>Place the host pupae into a Petri dish with damp filter paper and cotton balls, and incubate them at 25 &#176;C with roughly 70% humidity until injected embryos hatch (roughly 1-2 days) (Figure 2- vii). 
	NOTE: Importantly, hosts can be left with a peeled off puparium and the N. vitripennis eggs will develop normally so long as they are incubated in a humidified chamber (Petri dish with damp filter paper and cotton balls) to prevent desiccation. Incubating at room temperature and with humidity from the dampened cotton balls is sufficient in case humidity cannot be controlled.</li>
	<li>Transfer the host pupae into a new Petri dish at 70% humidity and 25&#176;C after G0 embryos hatch.
	<ol>
		<li>Monitor the host pupae and injected G0 larvae daily. If the host pupae have a foul smell, transfer the injected larvae to new healthy host pupae.</li>
	</ol>
	</li>
</ol>
6. Screening for Genome Modifications
<ol>
	<li>Remove each N. vitripennis pupae from the host using either a fine paintbrush or embryo pick. Injected G0 larvae should pupate around 8 days post injection.</li>
	<li>Place each removed pupa into an isolated glass vial plugged with cotton and allow them to emerge as G0 adults, ensuring that the hatched females will be virgin which will help with downstream genetic crosses. 
	NOTE: Females can be sorted from males by wing length: females have longer wings which extend past the body profile when turned on the side. All microinjected female pupae can be placed together into a plugged vial, and the same applies to male pupae.</li>
	<li>Screen G0 individuals, after eclosion, by either PCR or visually (if disrupting a phenotypic gene). For example, if CRISPR is used to create deletions in a given gene and the disrupted gene confers a visible phenotype, then sort and keep individuals with a mutant version of the affected phenotype (Figure 2- viii)21. If, however, the affected gene encodes for a non-visible phenotype, such as a mutant of an essential gene, perform a crossing scheme (cross G0 male with wild type female, or cross G0 female with wild type male) to recover and screen for heritable mutants through assaying lethality or sterility. 
	NOTE: Similar to D. melanogaster, N. vitripennis adults can be immobilized by exposure to CO2 allowing for straightforward manipulation. Moreover, unmated females will give rise to 100% haploid male broods, so therefore a mutant unmated female can give rise to large number of mutant males that can be used for subsequent analysis. 
	NOTE: Mutations in essential genes will need to be kept by mating surviving G0 injected females (presumably heterozygous for a mutation) to wild type males; in this particular case, half of the G1 male progeny should die due to inheritance of the lethal mutation, while half of the G1 female progeny will be carriers of the lethal.</li>
	<li>Outcross G0 adults and screen G1 progeny for transgene insertions using transgenesis markers if the goal is transgenesis. Currently transgenesis remains to be demonstrated in N. vitripennis.</li>
</ol>

<ol><li>Rivers, D. B., Denlinger, D. L. <a target="_blank" href="http://www.sciencedirect.com/science/article/pii/0022191094900833">Developmental fate of the flesh fly, Sarcophaga bullata, envenomated by the pupal ectoparasitoid, Nasonia vitripennis.</a> J Insect Physiol. 40 (2), 121-127 (1994).</li>
<li>Whiting, A. R. <a target="_blank" href="https://www.jstor.org/stable/2818420?seq=1#page_scan_tab_contents">The Biology of the Parasitic Wasp Mormoniella vitripennis [=Nasonia brevicornis] (Walker).</a> Q Rev Biol. 42 (3), 333-406 (1967).</li>
<li>Ferree, P. M., Avery, A., Azpurua, J., Wilkes, T., Werren, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+bacterium+targets+maternally+inherited+centrosomes+to+kill+males+in+Nasonia%5BTitle%5D)+AND+%22Curr+Biol%22%5BJournal%5D)">A bacterium targets maternally inherited centrosomes to kill males in Nasonia.</a> Curr Biol. 18 (18), 1409-1414 (2008).</li>
<li>Hurst, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Male-Killing+Bacteria+in+Insects%3A+Mechanisms%2C+Incidence%2C+and+Implications%5BTitle%5D)+AND+%22Emerg+Infect+Dis%22%5BJournal%5D)">Male-Killing Bacteria in Insects: Mechanisms, Incidence, and Implications.</a> Emerg Infect Dis. 6 (4), 329-336 (2000).</li>
<li>Aldrich, J. C., Leibholz, A., Cheema, M. S., Ausiό, J., Ferree, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+%26%2334%3Bselfish%26%2334%3B+B+chromosome+induces+genome+elimination+by+disrupting+the+histone+code+in+the+jewel+wasp+Nasonia+vitripennis%5BTitle%5D)+AND+%22Sci+Rep%22%5BJournal%5D)">A &#34;selfish&#34; B chromosome induces genome elimination by disrupting the histone code in the jewel wasp Nasonia vitripennis.</a> Sci Rep. 7, 42551(2017).</li>
<li>Werren, J. H., Stouthamer, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((PSR+%28paternal+sex+ratio%29+chromosomes%3A+the+ultimate+selfish+genetic+elements%5BTitle%5D)+AND+%22Genetica%22%5BJournal%5D)">PSR (paternal sex ratio) chromosomes: the ultimate selfish genetic elements.</a> Genetica. 117 (1), 85-101 (2003).</li>
<li>Nur, U., Werren, J. H., Eickbush, D. G., Burke, W. D., Eickbush, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+%26%2334%3Bselfish%26%2334%3B+B+chromosome+that+enhances+its+transmission+by+eliminating+the+paternal+genome%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">A &#34;selfish&#34; B chromosome that enhances its transmission by eliminating the paternal genome.</a> Science. 240 (4851), 512-514 (1988).</li>
<li>Danneels, E. L., Rivers, D. B., de Graaf, D. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Venom+Proteins+of+the+Parasitoid+Wasp+Nasonia+vitripennis%3A+Recent+Discovery+of+an+Untapped+Pharmacopee%5BTitle%5D)+AND+%22Toxins%22%5BJournal%5D)">Venom Proteins of the Parasitoid Wasp Nasonia vitripennis: Recent Discovery of an Untapped Pharmacopee.</a> Toxins. 2 (4), 494-516 (2010).</li>
<li>de Graaf, D. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Insights+into+the+venom+composition+of+the+ectoparasitoid+wasp+Nasonia+vitripennis+from+bioinformatic+and+proteomic+studies%5BTitle%5D)+AND+%22Insect+Mol+Biol%22%5BJournal%5D)">Insights into the venom composition of the ectoparasitoid wasp Nasonia vitripennis from bioinformatic and proteomic studies.</a> Insect Mol Biol. 19 Suppl 1, 11-26 (2010).</li>
<li>Verhulst, E. C., Beukeboom, L. W., van de Zande, L. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Maternal+control+of+haplodiploid+sex+determination+in+the+wasp+Nasonia%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Maternal control of haplodiploid sex determination in the wasp Nasonia.</a> Science. 328 (5978), 620-623 (2010).</li>
<li>Grillenberger, B. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Genetic+structure+of+natural+Nasonia+vitripennis+populations%3A+validating+assumptions+of+sex-ratio+theory%5BTitle%5D)+AND+%22Mol+Ecol%22%5BJournal%5D)">Genetic structure of natural Nasonia vitripennis populations: validating assumptions of sex-ratio theory.</a> Mol Ecol. 17 (12), 2854-2864 (2008).</li>
<li>Pultz, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+major+role+for+zygotic+hunchback+in+patterning+the+Nasonia+embryo%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">A major role for zygotic hunchback in patterning the Nasonia embryo.</a> Development. 132 (16), 3705-3715 (2005).</li>
<li>Lynch, J. A., Brent, A. E., Leaf, D. S., Pultz, M. A., Desplan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Localized+maternal+orthodenticle+patterns+anterior+and+posterior+in+the+long+germ+wasp+Nasonia%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Localized maternal orthodenticle patterns anterior and posterior in the long germ wasp Nasonia.</a> Nature. 439 (7077), 728-732 (2006).</li>
<li>Lynch, J. A., Olesnicky, E. C., Desplan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Regulation+and+function+of+tailless+in+the+long+germ+wasp+Nasonia+vitripennis%5BTitle%5D)+AND+%22Dev+Genes+Evol%22%5BJournal%5D)">Regulation and function of tailless in the long germ wasp Nasonia vitripennis.</a> Dev Genes Evol. 216 (7-8), 493-498 (2006).</li>
<li>Werren, J. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Functional+and+evolutionary+insights+from+the+genomes+of+three+parasitoid+Nasonia+species%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Functional and evolutionary insights from the genomes of three parasitoid Nasonia species.</a> Science. 327 (5963), 343-348 (2010).</li>
<li>Ferree, P. M., Fang, C., Mastrodimos, M., Hay, B. A., Amrhein, H., Akbari, O. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Identification+of+Genes+Uniquely+Expressed+in+the+Germ-Line+Tissues+of+the+Jewel+Wasp+Nasonia+vitripennis%5BTitle%5D)+AND+%22G3%22%5BJournal%5D)">Identification of Genes Uniquely Expressed in the Germ-Line Tissues of the Jewel Wasp Nasonia vitripennis.</a> G3. 5 (12), 2647-2653 (2015).</li>
<li>Akbari, O. S., Antoshechkin, I., Hay, B. A., Ferree, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Transcriptome+profiling+of+Nasonia+vitripennis+testis+reveals+novel+transcripts+expressed+from+the+selfish+B+chromosome%2C+paternal+sex+ratio%5BTitle%5D)+AND+%22G3%22%5BJournal%5D)">Transcriptome profiling of Nasonia vitripennis testis reveals novel transcripts expressed from the selfish B chromosome, paternal sex ratio.</a> G3. 3 (9), 1597-1605 (2013).</li>
<li>Sackton, T. B., Werren, J. H., Clark, A. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Characterizing+the+infection-induced+transcriptome+of+Nasonia+vitripennis+reveals+a+preponderance+of+taxonomically-restricted+immune+genes%5BTitle%5D)+AND+%22PloS+one%22%5BJournal%5D)">Characterizing the infection-induced transcriptome of Nasonia vitripennis reveals a preponderance of taxonomically-restricted immune genes.</a> PloS one. 8 (12), e83984(2013).</li>
<li>Werren, J. H., Loehlin, D. W., Giebel, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Larval+RNAi+in+Nasonia+%28parasitoid+wasp%29%5BTitle%5D)+AND+%22Cold+Spring+Harb+Protoc%22%5BJournal%5D)">Larval RNAi in Nasonia (parasitoid wasp).</a> Cold Spring Harb Protoc. (10), (2009).</li>
<li>Lynch, J. A., Desplan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+method+for+parental+RNA+interference+in+the+wasp+Nasonia+vitripennis%5BTitle%5D)+AND+%22Nat+Protoc%22%5BJournal%5D)">A method for parental RNA interference in the wasp Nasonia vitripennis.</a> Nat Protoc. 1 (1), 486-494 (2006).</li>
<li>Li, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Generation+of+heritable+germline+mutations+in+the+jewel+wasp+Nasonia+vitripennis+using+CRISPR%2FCas9%5BTitle%5D)+AND+%22Sci+Rep%22%5BJournal%5D)">Generation of heritable germline mutations in the jewel wasp Nasonia vitripennis using CRISPR/Cas9.</a> Sci Rep. 7 (1), 901(2017).</li>
<li>Meer, M. M. M., Witteveldt, J., Stouthamer, R. <a target="_blank" href="http://onlinelibrary.wiley.com/doi/10.1046/j.1570-7458.1999.00594.x/full">Development of a microinjection protocol for the parasitoid Nasonia vitripennis.</a> Entomol Exp Appl. 93 (3), 323-327 (1999).</li>
<li>Werren, J. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Functional+and+evolutionary+insights+from+the+genomes+of+three+parasitoid+Nasonia+species%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Functional and evolutionary insights from the genomes of three parasitoid Nasonia species.</a> Science. 327 (5963), 343-348 (2010).</li>
<li>Doudna, J. A., Charpentier, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+new+frontier+of+genome+engineering+with+CRISPR-Cas9%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">The new frontier of genome engineering with CRISPR-Cas9.</a> Science. 346 (6213), 1258096(2014).</li>
</ol>

The authors have nothing to disclose.

The recent sequencing of the N. vitripennis genome has unleashed an important need for molecular tools to functionally characterize unknown genes within this species23. The CRISPR-Cas9 system, and many other gene editing tools, have proven to be valuable in investigating gene functions for a number of organisms24. However, to generate heritable mutations, these tools require performing embryo microinjections. Therefore, demonstrated here is a detailed visual technique that includes a number of innovations that allow for efficient N. vitripennis embryo microinjections.
Overall, this detailed technique offers a number of significant innovations, with respect to existing methods23, that allow for efficient N. vitripennis embryo microinjections. For example, to facilitate the rapid collection of embryos, an oviposition tool (foam stopper) is created and used to restrict egg laying entirely to the posterior end of the host (Figure 1), which greatly facilitates collection of numerous embryos within a short period of time. Techniques for rearing wasp colonies with large numbers to collect greater numbers of eggs are also improved and defined. Additionally, to accelerate embryo alignment, an embryo alignment device which allows for embryos to be efficiently aligned and injected without having to use double-sided sticky tape to secure the embryos in place (Figure 3) is developed.
Furthermore, it is found that by keeping the embryos moist with water during injection and not desiccating the embryos or covering them with oil improved survival rates. Additionally, several capillary glass types are tested and parameters to construct the perfect needles for N. vitripennis microinjection (Table 1, Figure 4) are determined. Moreover, following microinjection, embryo survival rates are able to significantly increase due to incubating injected eggs in pre-stung hosts placed in high-humidity (70%) chambers. These innovations allow for a more streamlined and successful microinjection procedure for N. vitripennis.
Minor modifications in terms of injection apparatus, and rearing procedures can be made to this protocol, depending on user preferences. However, a number of critical steps will be essential for successfully generating mutants in N. vitripennis. For example, working quickly so that injected embryos are &#62; 1 h old, and ensuring that injection needles are sharp enough to minimize damage to the embryo will both be essential. While this protocol should be effective for generating mutations in many (if not all) genes, one major limitation is the CRIPSR/Cas9 requirement to target a PAM (NGG) sequence which will dictate the target sequence.
In conclusion, this improved technique can be used to generate many kinds of genome modifications, such as mutations, deletions, and possibly even insertions using CRIPSR/Cas9 technologies21, or even transgene insertions to generate transgenic N. vitripennis, which should greatly accelerate functional research in this organism.

The jewel wasp, N. vitripennis, is one of the four species within the genus Nasonia that are ectoparasitoids of flesh eating flies such as Sarcophaga bullata1. Due to their fast-generational periods, ease of rearing in the laboratory, and a range of unique and important biological attributes, N. vitripennis has been a focus for the development of multiple experimental tools found in traditional model organisms. For example, some unique biological attributes include a haplo-diploid reproduction system2, a relationship with microbial and genetic parasites3,4, and a supernumary (B) chromosome5,6,7. Taken together, these&#160;make N. vitripennis an important experimental system for experiments aimed at elucidating the molecular and cellular aspects of these processes, in addition to others including venom production8,9, sex determination10,11, and evolution and development of axis pattern formation12,13,14. Moreover, the genetic toolkit to study the biology of N. vitripennis has dramatically increased over the last decade or so, with the sequencing of a high-resolution genome15, several gene expression studies16,17,18, and the ability to functionally disrupt gene expression relying on RNA interference (RNAi)19,20, which together have improved the tractability and capabilities of performing reverse genetics in this organism.
Despite the many important scientific advancements and expanded toolkits in this organism, as of present knowledge only one group has successfully performed embryonic microinjections to generate heritable genome modifications21. This is primarily due to the difficulties of working with embryos of N. vitripennis as they are quite fragile and small, being ~2/3 the size of Drosophila melanogaster embryos, making them generally difficult to manipulate. Additionally, N. vitripennis females deposit their eggs into pre-stung blowfly pupae, within which the entirety of embryogenesis, larval, and pupal development occurs. Therefore, for successful microinjections, pre-blastoderm stage, embryos must be efficiently collected from host pupae, quickly microinjected, and immediately transplanted back into their hosts for development. These steps require precision and dexterity to avoid damaging the microinjected embryos, or the pupal hosts, making the technique exceptionally challenging. Notwithstanding, there is one short protocol published over a decade ago that describes N. vitripennis embryo microinjection22. However, this procedure requires that the freshly laid embryos be desiccated, it uses sticky tape to anchor the eggs for microinjection, and does not include a visual demonstration of the technique. Therefore, described here is an updated and revised protocol, including a visual procedure, detailing an improved step-by-step protocol for N. vitripennis embryo microinjections that can be followed by any basic lab to generate heritable genome modifications in this important model insect.

<table><tbody><tr><td>Bugdorm</td><td>Bugdorm</td><td>41515</td><td>Insect Rearing Cage</td></tr><tr><td>Glass Test Tubes</td><td>Fisher Scientific</td><td>982010</td><td></td></tr><tr><td>Flesh fly pupae, &lt;em&gt;Sarcophaga bullata&lt;/em&gt;</td><td>Carolina Insects</td><td>144440</td><td></td></tr><tr><td>Microelectrode Beveler</td><td>Sutter Instruments</td><td>BV10</td><td></td></tr><tr><td>Diamond abrasive plate (0.7u to 2.0u tip sizes)</td><td>Sutter Instruments</td><td>104E</td><td></td></tr><tr><td>Micromanipulator</td><td>World Precision Instruments</td><td>Kite R</td><td></td></tr><tr><td>Femtojet Express programmable microinjector</td><td>Eppendorf</td><td></td><td></td></tr><tr><td>Micropipette Puller</td><td>Sutter Instruments</td><td>P-1000 or P-2000</td><td></td></tr><tr><td>Stereo Microscope</td><td>Olympus</td><td>SZ51</td><td></td></tr><tr><td>Compound Microscope</td><td>Leica DM-750</td><td></td><td></td></tr><tr><td>Aluminosilicate glass capillary tubing 1mm(outside diameter) X 0.58mm (inner diameter)</td><td>Sutter Instruments</td><td>BF100-58-10</td><td>Can also use Borosilicate or Quartz</td></tr><tr><td>Identi-Plugs</td><td>JAECE</td><td>L800-B</td><td>Foam plugs</td></tr><tr><td>Microscope Slides</td><td>Fisherbrand</td><td>12-550-A3</td><td></td></tr><tr><td>Micro Cover glass</td><td>VWR</td><td>48366045</td><td></td></tr><tr><td>Adhesive</td><td>Aron Alpha</td><td>AA471</td><td>For glueing coverslip onto microscope slide</td></tr><tr><td>Fine-tip paintbrush</td><td>ZEM</td><td>2595</td><td></td></tr><tr><td>Ultra-fine tip forcep</td><td>Fisher Scientific</td><td>16-100-121</td><td></td></tr><tr><td>Femtotips Microloader tips</td><td>Fisher Scientific</td><td>E5242956003</td><td></td></tr></tbody></table>

embryo microinjection and transplantation technique for nasonia vitripennis genome manipulation

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex determination, venom synthesis, and host-symbiont interactions, among others. A major limitation of working with this organism is the lack of effective protocols to perform directed genome modifications. An important part of genome modification is delivery of editing reagents, including CRISPR/Cas9 molecules, into embryos through microinjection. While microinjection is well established in many model organisms, this technique is particularly challenging to perform in N. vitripennis primarily due to its small embryo size, and the fact that embryonic development occurs entirely within a parasitized blowfly pupa. The following procedure overcomes these significant challenges while demonstrating a streamlined, visual procedure for effectively removing wasp embryos from parasitized host pupae, microinjecting them, and carefully transplanting them back into the host for continuation and completion of development. This protocol will strongly enhance the capability of research groups to perform advanced genome modifications in this organism.

This protocol provides detailed guidelines for colony rearing, pre-blastoderm embryo collection, alignment, microinjection, and subsequent transplantation after injection and can be used for efficient genome engineering in N. vitripennis. As shown in Figure 2, the general sequence of steps for a successful microinjection into N. vitripennis include: (i) permitting male and female adults to mate (~ 4 days), (ii) supplying fresh host fly pupae (S. bullata) placed in a modified foam plug to mated females and allowing for oviposition (~ 45 min), (iii) carefully peeling away the parasitized host pupae cuticle to expose and collect pre-blastoderm stage wasp embryos (~ 15 min), (iv) aligning collected embryos (~ 15 min), (v) microinjecting genome modification components into embryos (~ 15 min), (vi) carefully placing injected embryos back into pre-stung hosts to allow for proper development (~ 15 min), and (vii) preventing dehydration of the injected embryo/host by transferring them into a humidified chamber with roughly 70% relative humidity (~ 15 min). Parasitized hosts are then incubated for roughly 14 days to allow for complete development of N. vitripennis embryos. Once injected, adults emerge from the host (viii), isolate, mate, and screen them individually for the presence of expected mutations.
For effective needle penetration and microinjection into N. vitripennis embryos, several types of capillary glass needles with filament including quartz, aluminosilicate, and borosilicate types are tested. It is found that the quality of needle is critical for avoiding breakage/clogging during injection, and for achieving high rates of both embryo survival and transformation efficiency. For each glass type, an effective protocol was developed to pull needles in order to have a desired hypodermic-like long tip effective for N. vitripennis embryo microinjection using different micropipette pullers (P-1000, and P-2000) (Table 1, Figure 4).
To optimize the procedure, survival rates are measured following injection of varying amounts of genome modification components. The genome modification components used here were mixed guide RNAs and Cas9 protein for CRISPR-mediated genome editing, which were demonstrated previously to work well in N. vitripennis21. Similar to what was previously reported, here a sgRNA targeting the cinnabar gene is designed and synthesized. By targeting and disrupting this gene, an easily identifiable phenotypic change is seen in the eye color of the organism19,21. An injection mixture combining a variety of concentrations of sgRNA (0, 20, 40, 80, 160, and 320 ng/&#181;L) with Cas9 protein (0, 20, 40, 80, 160, and 320 ng/&#181;L) is created and injected into embryos of wild type N. vitripennis. Survival rate of injected embryos is found to be dose-dependent (Table 2)21. The increased concentration of sgRNA and Cas9 protein lead to decreased survival rates (Table 2), perhaps due to additive off-target effects. High humidity (~ 70%) is also found to be important for embryo survival after transplantation to hosts, as low humidity (~ 10%) resulted in 100% death to all injected embryos.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56990/56990fig1.jpg" /> 
Figure 1. Preparation of host pupae (S. bullata) for N. vitripennis embryo oviposition. (A)Young and old S. bullata pupae. Older pupae have a darker cuticle whereas younger pupae have a more reddish tint to their cuticle. Younger pupae are preferred for maximizing oviposition. Posterior and anterior ends of the pupae can also be distinguished by a &#34;crater-like opening on the posterior end, whereas the anterior end comes to a rounded point. (B) Host (S. bullata) pupa preparation for N. vitripennis embryo oviposition. Inserting the host pupae into a foam plug that has had a pupae sized hole carved out. Have the posterior side of the host pupae face inside the plug while 0.2 cm of the anterior end exposed to allow for maximum concentration of oviposition into the anterior area. <a href="https://cloudflare.jove.com/files/ftp_upload/56990/56990fig1large.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/56990/56990fig2.jpg" /> 
Figure 2. Timeline for creating N. vitripennis mutants by microinjection. Timeline of N. vitripennis embryo collection, CRISPR/Cas9 microinjection, and post-injection procedures. N. vitripennis adults were allowed to mate in absence of an oviposition site for 4 days (i). Following, a fresh, flesh fly host pupae, S. bullata, placed inside a foam stopper as to only expose 0.5 cm of the posterior end, was introduced to the gravid females for 45 min to allow for parasitization (ii). Concurrently injection materials including microinjection needles and CRISPR/Cas9 components were prepared (iii). Embryos were collected from the host (iv), aligned (v), and injected with CRISPR/Cas9 components (vi). The injected embryos were carefully transferred back to a pre-stung host (vii) and incubated till fully developed (14 days) (viii). When the adults emerged, mutants were screened for phenotypes of expected CRISPR/Cas9 induced mutations in the target gene (ix). The entire procedure generally takes 19 days to complete. <a href="https://cloudflare.jove.com/files/ftp_upload/56990/56990fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56990/56990fig3.jpg" /> 
Figure 3. Modified microscope slide for lining embryos. (A) A coverslip. (B) A microscope slide. (C) An embryo alignment device. A coverslip can be glued onto a microscope slide to be used to line embryos for injection. The purpose of the coverslip is to act as an edge to allow for easy manipulation and lining of embryos. <a href="https://cloudflare.jove.com/files/ftp_upload/56990/56990fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56990/56990fig4.jpg" /> 
Figure 4. Injection needle preparation. (A) Examples of good and bad aluminosilicate glass needles. (B) The tips of an unbeveled, correctly beveled, and poorly beveled needle. Aluminosilicate glass capillary tubes were pulled by using a micropipette puller. The produced needle tips were then gently opened and refined using a beveler. The good bevelled needle has a very sharp tip, and the bad bevelled needle has a blunt tip. <a href="https://cloudflare.jove.com/files/ftp_upload/56990/56990fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Capillary Glass Type</td>
			<td>Sutter Needle Puller Model</td>
			<td>Heat</td>
			<td>Filament</td>
			<td>Velocity</td>
			<td>Delay</td>
			<td>Pull</td>
			<td>Pressure</td>
		</tr>
		<tr>
			<td>Quartz</td>
			<td>P-2000</td>
			<td>750</td>
			<td>4</td>
			<td>40</td>
			<td>150</td>
			<td>165</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Aluminosilicate</td>
			<td>P-1000</td>
			<td>605</td>
			<td>-</td>
			<td>130</td>
			<td>80</td>
			<td>70</td>
			<td>500</td>
		</tr>
		<tr>
			<td>Borosilicate</td>
			<td>P-1000</td>
			<td>450</td>
			<td>-</td>
			<td>130</td>
			<td>80</td>
			<td>70</td>
			<td>500</td>
		</tr>
	</tbody>
</table>
Table 1. Settings for needle puller. 
Note: Needle puller setting vary from machine to machine so each lab will need to optimize their own needle puller settings. This table has been modified from Li et al.21
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="3">sgRNA-1</td>
			<td rowspan="3">Cas9</td>
			<td rowspan="3">Total embryos</td>
			<td>Transplantation (10% humidity)</td>
			<td colspan="4">Transplantation (70% humidity)</td>
		</tr>
		<tr>
			<td>Larvae Survivors</td>
			<td>Larvae Survivors</td>
			<td colspan="3">Adult survivors</td>
		</tr>
		<tr>
			<td>Total (%)</td>
			<td>Total (%)</td>
			<td>&#9794;</td>
			<td>&#9792;</td>
			<td>Total (%)</td>
		</tr>
		<tr>
			<td>No injection</td>
			<td>No injection</td>
			<td>100</td>
			<td>94 (94)</td>
			<td>96 (96)</td>
			<td>66</td>
			<td>26</td>
			<td>92 (92)</td>
		</tr>
		<tr>
			<td>Water</td>
			<td>Water</td>
			<td>100</td>
			<td>0 (0)</td>
			<td>78 (78)</td>
			<td>44</td>
			<td>32</td>
			<td>76 (76)</td>
		</tr>
		<tr>
			<td>20 ng/&#181;L</td>
			<td>20 ng/&#181;L</td>
			<td>100</td>
			<td>0 (0)</td>
			<td>74 (74)</td>
			<td>34</td>
			<td>34</td>
			<td>68 (68)</td>
		</tr>
		<tr>
			<td>40 ng/&#181;L</td>
			<td>40 ng/&#181;L</td>
			<td>100</td>
			<td>0 (0)</td>
			<td>67 (67)</td>
			<td>30</td>
			<td>32</td>
			<td>62 (62)</td>
		</tr>
		<tr>
			<td>80 ng/&#181;L</td>
			<td>80 ng/&#181;L</td>
			<td>100</td>
			<td>0 (0)</td>
			<td>53 (53)</td>
			<td>24</td>
			<td>22</td>
			<td>46 (46)</td>
		</tr>
		<tr>
			<td>160 ng/&#181;L</td>
			<td>160 ng/&#181;L</td>
			<td>100</td>
			<td>0 (0)</td>
			<td>41 (41)</td>
			<td>16</td>
			<td>22</td>
			<td>38 (38)</td>
		</tr>
		<tr>
			<td>320 ng/&#181;L</td>
			<td>320 ng/&#181;L</td>
			<td>100</td>
			<td>0 (0)</td>
			<td>25 (25)</td>
			<td>10</td>
			<td>10</td>
			<td>20 (20)</td>
		</tr>
	</tbody>
</table>
Table 2. Injection and transplantation survivorship and mutagenesis rates based on injections of different concentrations of sgRNA and Cas9. This table has been modified from Li et al.21

Watch this Scientific Journal Video about Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation at JoVE.com

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Microinjection of Nasonia vitripennis embryos is an essential method for generating heritable genome modifications. Described here is a detailed procedure for microinjection and transplantation of Nasonia vitripennis embryos, which will greatly facilitate future genome manipulation in this organism.

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex ...

embryo-microinjection-transplantation-technique-for-nasonia

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Genetics

12.2K Views.  University of California, Riverside. Microinjection van Nasonia vitripennis embryo's is een essentiële methode voor het genereren van de wijzigingen van het erfelijk genoom. Hier beschreven is een gedetailleerde procedure voor microinjection en transplanteren van Nasonia vitripennis embryo's, die toekomstige genoom manipulatie in dit organisme aanzienlijk zal vergemakkelijken.

Video: Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. This is due to its reproductive behavior: females damage the external surface of fruits and vegetables when they lay eggs and the hatched larvae feed on their pulp. Wild C. capitata populations are traditionally controlled through insecticide spraying and/or eco-friendly approaches, the most successful being the Sterile Insect Technique (SIT). The SIT relies on mass-rearing, radiation-based sterilization and field release of males that retain their capacity to mate but are not able to generate fertile progeny. The advent and the subsequent rapid development of biotechnological tools, together with the availability of the medfly genome sequence, has greatly boosted our understanding of the biology of this species. This favored the proliferation of new strategies for genome manipulation, which can be applied to population control.
In this context, embryo microinjection plays a dual role in expanding the toolbox for medfly control. The ability to interfere with the function of genes that regulate key biological processes, indeed, expands our understanding of the molecular machinery underlying medfly invasiveness. Furthermore, the ability to achieve germ-line transformation facilitates the production of multiple transgenic strains that can be tested for future field applications in novel SIT settings. Indeed, genetic manipulation can be used to confer desirable traits that can, for example, be used to monitor sterile male performance in the field, or that can result in early life-stage lethality. Here we describe a method to microinject nucleic acids into medfly embryos to achieve these two main goals.

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Diptera: Tephritidae) is a worldwide pest of agriculture. A deeper understanding of its biology is key to control medfly populations and thus reduce economic impact. Embryo microinjection is a fundamental tool allowing both germ-line transformation and reverse genetics studies in this species.

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. ...

Loss- and Gain-of-function Approach to Investigate Early Cell Fate Determinants in Preimplantation Mouse Embryos

Gene silencing and overexpression techniques are instrumental for the identification of genes involved in embryonic development. Direct target gene modification in preimplantation embryos provides a means to study the underlying mechanisms of genes implicated in, for instance, cellular differentiation into the trophectoderm (TE) and the inner cell mass (ICM). Here, we describe a protocol that examines the role of neogenin as an authentic receptor for initial cell fate determination in preimplantation mouse embryos. First, we discuss the experimental manipulations that were used to produce gain and loss of neogenin function by microinjecting neogenin cDNA and shRNA; the effectiveness of this approach was confirmed by a strong correlation between the pair-wise expression levels of either red fluorescent protein (RFP) or green fluorescent protein (GFP) and the immunocytochemical quantification of neogenin expression. Secondly, overexpression of neogenin in preimplantation mouse embryos leads to normal ICM development while neogenin knockdown causes the ICM to develop abnormally, implying that neogenin could be a receptor that relays extracellular cues to drive blastomeres to early cell fates. Given the success of this detailed protocol in investigating the function of a novel embryonic developmental stage-specific receptor, we propose that it has the potential to aid in exploration and identification of other stage-specific genes during embryogenesis.

The goal of this protocol is to describe a loss- and gain-of function method that is applicable to identify neogenin as a stage-specific receptor that leads to trophectoderm and inner cell mass differentiation in preimplantation mouse embryos.

 Gene silencing and overexpression techniques are instrumental for the identification of genes involved in embryonic development. Direct target gene ...

Developmental Biology

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance to man of conventional invertebrate model systems and the complexity of genomes in vertebrates are critical challenges to modeling human normal and pathological conditions. The chordate Ciona intestinalis is an invertebrate chordate that emerged from a common ancestor with the vertebrates and may represent features at the interface between invertebrates and vertebrates. A common body plan with much simpler cellular and genomic composition should unveil gene regulatory network (GRN) links and functional genomics readouts explaining phenomena in the vertebrate condition. The compact genome of Ciona, a fixed embryonic lineage with few divisions and large cells, combined with versatile community tools foster efficient gene functional analyses in this organism. Here, we present several crucial methods for this promising model organism, which belongs to the closest sister group to vertebrates. We present protocols for transient transgenesis by electroporation, along with microinjection-mediated gene knockdown, which together provide the means to study gene function and genomic regulatory elements. We extend our protocols to provide information on how community databases are utilized for in silico design of gene regulatory or gene functional experiments. An example study demonstrates how novel information can be gained on the interplay, and its quantification, of selected neural factors conserved between Ciona and man. Furthermore, we show examples of differential subcellular localization in embryonic cells, following DNA electroporation in Ciona zygotes. Finally, we discuss the potential of these protocols to be adapted for tissue specific gene interference with emerging gene editing methods. The in vivo approaches in Ciona overcome major shortcomings of classical model organisms in the quest of unraveling conserved mechanisms in the chordate developmental program, relevant to stem cell research, drug discovery, and subsequent clinical application.

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

We present transient transgenesis and gene knockdown in Ciona intestinalis, a chordate sister group to vertebrates, using microinjection and electroporation techniques. Such methods facilitate functional genomics in this simple invertebrate that features rudimentary characteristics of vertebrates, including notochord and head sensory epithelia, and many orthologs of human disease associated genes.

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance ...

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, Eastern equine encephalitis, lymphatic filariasis, and Saint Louis encephalitis. Notably, avian malaria has played a major role in the extinction of numerous endemic island bird species, while WNV has become an important vector-borne disease in the United States. To gain further insight into C. quinquefasciatus biology and expand their genetic control toolbox, we need to develop more efficient and affordable methods for genome engineering in this species. However, some biological traits unique to Culex mosquitoes, particularly their egg rafts, have made it difficult to perform microinjection procedures required for genome engineering. To address these challenges, we have developed an optimized embryo microinjection protocol that focuses on mitigating the technical obstacles associated with the unique characteristics of Culex mosquitoes. These procedures demonstrate optimized methods for egg collection, egg raft separation and other handling procedures essential for successful microinjection in C. quinquefasciatus. When coupled with the CRISPR/Cas9 genome editing technology, these procedures allow us to achieve site-specific, efficient and heritable germline mutations, which are required to perform advanced genome engineering and develop genetic control technologies in this important, but currently understudied, disease vector.

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

This protocol describes microinjection procedures for Culex quinquefasciatus embryos that are optimized to work with CRISPR/Cas9 gene editing tools. This technique can efficiently generate site-specific, heritable, germline mutations that can be used for building genetic technologies in this understudied disease vector.

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, ...

phiC31 Integrase-Mediated Embryo Transgenesis in Drosophila

Embryo Microinjection for Transgenesis in Drosophila

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the construction of transgenic flies. Microinjection requires some types of equipment, including a microinjector, a micromanipulator, an inverted microscope, and a stereo microscope.&#160;Plasmids isolated with a plasmid miniprep kit are qualified for microinjection. Embryos at the pre-blastoderm or syncytial blastoderm stage, where nuclei share a common cytoplasm, are subjected to microinjection. A cell strainer eases the process of dechorionating embryos. The optimal time for dechorionation and desiccation of embryos needs to be determined experimentally. To increase the efficiency of embryo microinjection, needles prepared by a puller need to be beveled by a needle grinder. In the process of grinding needles, we utilize a foot air pump with a pressure gauge to avoid the capillary effect of the needle tip. We routinely inject 120-140 embryos for each plasmid and obtain at least one transgenic line for around 85% of plasmids. This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and&#160;presents a detailed protocol for embryo microinjection for transgenesis in Drosophila.

Author Spotlight: Optimizing Embryo Microinjection for Transgenesis in Drosophila

This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and presents an optimized protocol for embryo microinjection,&#160;a crucial step for creating transgenic flies.

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the ...

Microinjection of Western Corn Rootworm, Diabrotica virgifera virgifera, Embryos for Germline Transformation, or CRISPR/Cas9 Genome Editing

The western corn rootworm (WCR) is an important pest of corn and is well known for its ability to rapidly adapt to pest management strategies. Although RNA interference (RNAi) has proved to be a powerful tool for studying WCR biology, it has its limitations. Specifically, RNAi itself is transient (i.e. does not result in long-term Mendelian inheritance of the associated phenotype), and it requires knowing the DNA sequence of the target gene. The latter can be limiting if the phenotype of interest is controlled by poorly conserved, or even novel genes, because identifying useful targets would be challenging, if not impossible. Therefore, the number of tools in WCR&#39;s genomic toolbox should be expanded by the development of methods that could be used to create stable mutant strains and enable sequence-independent surveys of the WCR genome. Herein, we detail the methods used to collect and microinject precellular WCR embryos with nucleic acids. While the protocols described herein are aimed at the creation of transgenic WCR, CRISPR/Cas9-genome editing could also be performed using the same protocols, with the only difference being the composition of the solution injected into the embryos.

Microinjection of Western Corn Rootworm, Diabrotica virgifera virgifera, Embryos for Germline Transformation, or CRISPR/Cas9 Genome Editing

Here we present protocols for collecting and microinjecting precellular western corn rootworm embryos for the purpose of performing functional-genomic assays such as germline transformation and CRISPR/Cas9-genome editing.

The western corn rootworm (WCR) is an important pest of corn and is well known for its ability to rapidly adapt to pest management strategies. Although ...

Bioengineering

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 ...

Biology

Microinjection of Corn Planthopper, Peregrinus maidis, Embryos for CRISPR/Cas9 Genome Editing

The corn planthopper, Peregrinus maidis, is&#160;a pest of maize and a vector of several maize viruses. Previously published methods describe the triggering of RNA interference (RNAi) in P. maidis through microinjection of double-stranded RNAs (dsRNAs) into nymphs and adults. Despite the power of RNAi, phenotypes generated via this technique are transient and lack long-term Mendelian inheritance. Therefore, the P. maidis toolbox needs to be expanded to include functional genomic tools that would enable the production of stable mutant strains, opening the door for researchers to bring new control methods to bear on this economically important pest. However, unlike the dsRNAs used for RNAi, the components used in CRISPR/Cas9-based genome editing and germline transformation do not easily cross cell membranes. As a result, plasmid DNAs, RNAs, and/or proteins must be microinjected into embryos before the embryo cellularizes, making the timing of injection a critical factor for success. To that end, an agarose-based egg-lay method was developed to allow embryos to be harvested from P. maidis females at relatively short intervals. Herein are provided detailed protocols for collecting and microinjecting precellular P. maidis embryos with CRISPR components (Cas9 nuclease that has been complexed with guide RNAs), and results of Cas9-based gene knockout of a P. maidis eye-color gene, white, are presented. Although these protocols describe CRISPR/Cas9-genome editing in P. maidis, they can also be used for producing transgenic P. maidis via germline transformation by simply changing the composition of the injection solution.

Microinjection of Corn Planthopper, Peregrinus maidis, Embryos for CRISPR/Cas9 Genome Editing

Herein are protocols for collecting and microinjecting precellular corn planthopper embryos for the purpose of modifying their genome via CRISPR/Cas9-based genome editing or for the addition of marked transposable elements through germline transformation.

The corn planthopper, Peregrinus maidis, is&#160;a pest of maize and a vector of several maize viruses. Previously published methods describe the ...

Mouse Genome Engineering Using Designer Nucleases

Transgenic mice carrying site-specific genome modifications (knockout, knock-in) are of vital importance for dissecting complex biological systems as well as for modeling human diseases and testing therapeutic strategies. Recent advances in the use of designer nucleases such as zinc finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and the clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system for site-specific genome engineering open the possibility to perform rapid targeted genome modification in virtually any laboratory species without the need to rely on embryonic stem (ES) cell technology. A genome editing experiment typically starts with identification of designer nuclease target sites within a gene of interest followed by construction of custom DNA-binding domains to direct nuclease activity to the investigator-defined genomic locus. Designer nuclease plasmids are in vitro transcribed to generate mRNA for microinjection of fertilized mouse oocytes. Here, we provide a protocol for achieving targeted genome modification by direct injection of TALEN mRNA into fertilized mouse oocytes.

Designer nucleases such as zinc finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs) can be used to modify the genome of mouse preimplantation embryos by triggering both the nonhomologous end joining (NHEJ) and homologous recombination (HR) pathways. These advances enable the rapid generation of mice with precise genetic modifications.

Transgenic mice carrying site-specific genome modifications (knockout, knock-in) are of vital importance for dissecting complex biological systems as well ...

A Method for Microinjection of Patiria minata Zygotes

Echinoderms have long been a favorite model system for studies of reproduction and development, and more recently for the study of gene regulation and evolution of developmental processes. The sea star, Patiria miniata, is gaining prevalence as a model system for these types of studies which were previously performed almost exclusively in the sea urchins, Strongylocentrotus purpuratus and Lytechinus variegatus. An advantage of these model systems is the ease of producing modified embryos in which a particular gene is up or downregulated, labeling a group of cells, or introducing a reporter gene. A single microinjection method is capable of creating a wide variety of such modified embryos. Here, we present a method for obtaining gametes from P. miniata, producing zygotes, and introducing perturbing reagents via microinjection. Healthy morphant embryos are subsequently isolated for quantitative and qualitative studies of gene function. The availability of genome and transcriptome data for this organism has increased the types of studies that are performed and the ease of executing them.

A Method for Microinjection of Patiria minata Zygotes

Methods that produce morphant embryos are essential to study developmental mechanisms and gene regulatory networks. The sea star Patiria miniata is an emerging model system for these studies. Here we present a protocol for obtaining gametes, producing cultures of embryos, and rapid microinjection of zygotes from this species.

Echinoderms have long been a favorite model system for studies of reproduction and development, and more recently for the study of gene regulation and ...

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

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