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>

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A., Desplan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Localized+maternal+orthodenticle+patterns+anterior+and+posterior+in+the+long+germ+wasp+Nasonia.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+and+function+of+tailless+in+the+long+germ+wasp+Nasonia+vitripennis.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+and+evolutionary+insights+from+the+genomes+of+three+parasitoid+Nasonia+species.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+Genes+Uniquely+Expressed+in+the+Germ-Line+Tissues+of+the+Jewel+Wasp+Nasonia+vitripennis.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transcriptome+profiling+of+Nasonia+vitripennis+testis+reveals+novel+transcripts+expressed+from+the+selfish+B+chromosome,+paternal+sex+ratio.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterizing+the+infection-induced+transcriptome+of+Nasonia+vitripennis+reveals+a+preponderance+of+taxonomically-restricted+immune+genes.">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. 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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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, Sarcophaga bullata</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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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 ...

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12.2K Views.  University of California, Riverside. 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.

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. ...

Single-cell Gene Expression Profiling Using FACS and qPCR with Internal Standards

Gene expression measurements from bulk populations of cells can obscure the considerable transcriptomic variation of individual cells within those populations. Single-cell gene expression measurements can help assess the role of noise in gene expression, identify correlations in the expression of pairs of genes, and reveal subpopulations of cells that respond differently to a stimulus. Here, we describe a procedure to measure the expression of up to 96 genes in single mammalian cells isolated from a population growing in tissue culture. Cells are sorted into lysis buffer by fluorescence-activated cell sorting (FACS), and the mRNA species of interest are reverse-transcribed and amplified. Gene expression is then measured using a microfluidic real-time PCR machine, which performs up to 96 qPCR assays on up to 96 samples at a time. We also describe the generation and use of PCR amplicon standards to enable the estimation of the absolute number of each transcript. Compared with other methods of measuring gene expression in single cells, this approach allows for the quantification of more distinct transcripts than RNA FISH at a lower cost than RNA-Seq.

We describe a method to sort single mammalian cells and to quantify the expression of up to 96 target genes of interest in each cell. This method includes the use of internal qPCR standards to enable the estimation of absolute transcript counts.

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Mapping RNA-RNA Interactions Globally Using Biotinylated Psoralen

Knowing how RNAs interact with themselves and with others is key to understanding RNA based gene regulation in the cell. While examples of RNA-RNA interactions such as microRNA-mRNA interactions have been shown to regulate gene expression, the full extent to which RNA interactions occur in the cell is still unknown. Previous methods to study RNA interactions have primarily focused on subsets of RNAs that are interacting with a particular protein or RNA species. Here, we detail a method named Sequencing of Psoralen crosslinked, Ligated, and Selected Hybrids (SPLASH) that allows genome-wide capture of RNA interactions in vivo in an unbiased manner. SPLASH utilizes in vivo crosslinking, proximity ligation, and high throughput sequencing to identify intramolecular and intermolecular RNA base-pairing partners globally. SPLASH can be applied to different organisms including bacteria, yeast and human cells, as well as diverse cellular conditions to facilitate the understanding of the dynamics of RNA organization under diverse cellular contexts. The entire experimental SPLASH protocol takes about 5 days to complete and the computational workflow takes about 7 days to complete.

Here, we detail the method of Sequencing of Psoralen crosslinked, Ligated, and Selected Hybrids (SPLASH), which enables genome-wide mapping of intramolecular and intermolecular RNA-RNA interactions in vivo. SPLASH can be applied to study RNA interactomes of organisms including yeast, bacteria and humans.

Knowing how RNAs interact with themselves and with others is key to understanding RNA based gene regulation in the cell. While examples of RNA-RNA ...

G2-seq: A High Throughput Sequencing-based Technique for Identifying Late Replicating Regions of the Genome

Numerous techniques have been developed to follow the progress of DNA replication through the S phase of the cell cycle. Most of these techniques have been directed toward elucidation of the location and timing of initiation of genome duplication rather than its completion. However, it is critical that we understand regions of the genome that are last to complete replication, because these regions suffer elevated levels of chromosomal breakage and mutation, and they have been associated with both disease and aging. Here we describe how we have extended a technique that has been used to monitor replication initiation to instead identify those regions of the genome last to complete replication. This approach is based on a combination of flow cytometry and high throughput sequencing. Although this report focuses on the application of this technique to yeast, the approach can be used with any cells that can be sorted in a flow cytometer according to DNA content.

We describe a technique for combining flow cytometry and high throughput sequencing to identify late replicating regions of the genome.

Numerous techniques have been developed to follow the progress of DNA replication through the S phase of the cell cycle. Most of these techniques have ...

Genome-wide Surveillance of Transcription Errors in Eukaryotic Organisms

Accurate transcription is required for the faithful expression of genetic information. Surprisingly though, little is known about the mechanisms that control the fidelity of transcription. To fill this gap in scientific knowledge, we recently optimized the circle-sequencing assay to detect transcription errors throughout the transcriptome of Saccharomyces cerevisiae, Drosophila melanogaster, and Caenorhabditis elegans. This protocol will provide researchers with a powerful new tool to map the landscape of transcription errors in eukaryotic cells so that the mechanisms that control the fidelity of transcription can be elucidated in unprecedented detail.

This protocol provides researchers with a new tool to monitor the fidelity of transcription in multiple model organisms.

Accurate transcription is required for the faithful expression of genetic information. Surprisingly though, little is known about the mechanisms that ...

Genome Editing in Mammalian Cell Lines using CRISPR-Cas

The clustered regularly interspaced short palindromic repeats (CRISPR) system functions naturally in bacterial adaptive immunity, but has been successfully repurposed for genome engineering in many different living organisms. Most commonly, the wildtype CRISPR associated 9 (Cas9) or Cas12a endonuclease is used to cleave specific sites in the genome, after which the DNA double-stranded break is repaired via the non-homologous end joining (NHEJ) pathway or the homology-directed repair (HDR) pathway depending on whether a donor template is absent or present respectively. To date, CRISPR systems from different bacterial species have been shown to be capable of performing genome editing in mammalian cells. However, despite the apparent simplicity of the technology, multiple design parameters need to be considered, which often leave users perplexed about how best to carry out their genome editing experiments. Here, we describe a complete workflow from experimental design to identification of cell clones that carry desired DNA modifications, with the goal of facilitating successful execution of genome editing experiments in mammalian cell lines. We highlight key considerations for users to take note of, including the choice of CRISPR system, the spacer length, and the design of a single-stranded oligodeoxynucleotide (ssODN) donor template. We envision that this workflow will be useful for gene knockout studies, disease modeling efforts, or the generation of reporter cell lines.

CRISPR-Cas is a powerful technology to engineer the complex genomes of plants and animals. Here, we detail a protocol to efficiently edit the human genome using different Cas endonucleases. We highlight important considerations and design parameters to optimize editing efficiency.

The clustered regularly interspaced short palindromic repeats (CRISPR) system functions naturally in bacterial adaptive immunity, but has been ...

Transforming, Genome Editing and Phenotyping the Nitrogen-fixing Tropical Cannabaceae Tree Parasponia andersonii

Parasponia andersonii is a fast-growing tropical tree that belongs to the Cannabis family (Cannabaceae). Together with 4 additional species, it forms the only known non-legume lineage able to establish a nitrogen-fixing nodule symbiosis with rhizobium. Comparative studies between legumes and P. andersonii could provide valuable insight into the genetic networks underlying root nodule formation. To facilitate comparative studies, we recently sequenced the P. andersonii genome and established Agrobacterium tumefaciens-mediated stable transformation and CRISPR/Cas9-based genome editing. Here, we provide a detailed description of the transformation and genome editing procedures developed for P. andersonii. In addition, we describe procedures for the seed germination and characterization of symbiotic phenotypes. Using this protocol, stable transgenic mutant lines can be generated in a period of 2-3 months. Vegetative in vitro propagation of T0 transgenic lines allows phenotyping experiments to be initiated at 4 months after A. tumefaciens co-cultivation. Therefore, this protocol takes only marginally longer than the transient Agrobacterium rhizogenes-based root transformation method available for P. andersonii, though offers several clear advantages. Together, the procedures described here permit P. andersonii to be used as a research model for studies aimed at understanding symbiotic associations as well as potentially other aspects of the biology of this tropical tree.

Transforming, Genome Editing and Phenotyping the Nitrogen-fixing Tropical Cannabaceae Tree Parasponia andersonii

Parasponia andersonii is a fast-growing tropical tree that belongs to the Cannabis family (Cannabaceae) and can form nitrogen-fixing root nodules in association with the rhizobium. Here, we describe a detailed protocol for reverse genetic analyses in P. andersonii based on Agrobacterium tumefaciens-mediated stable transformation and CRISPR/Cas9-based genome editing.

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Validating Whole Genome Nanopore Sequencing, using Usutu Virus as an Example

Whole genome sequencing can be used to characterize and to trace viral outbreaks. Nanopore-based whole genome sequencing protocols have been described for several different viruses. These approaches utilize an overlapping amplicon-based approach which can be used to target a specific virus or group of genetically related viruses. In addition to confirmation of the virus presence, sequencing can be used for genomic epidemiology studies, to track viruses and unravel origins, reservoirs and modes of transmission. For such applications, it is crucial to understand possible effects of the error rate associated with the platform used. Routine application in clinical and public health settings require that this is documented with every important change in the protocol. Previously, a protocol for whole genome Usutu virus sequencing on the nanopore sequencing platform was validated (R9.4 flowcell) by direct comparison to Illumina sequencing. Here, we describe the method used to determine the required read coverage, using the comparison between the R10 flow cell and Illumina sequencing as an example.

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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

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

Sand Fly (Phlebotomus papatasi) Embryo Microinjection for CRISPR/Cas9 Mutagenesis

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to visceral pathology. Deciphering the nature of the vector/parasite interactions is of primary importance for better understanding of Leishmania transmission to their hosts. Among the parameters controlling the sand fly vector competence (i.e. their ability to carry and transmit pathogens), parameters intrinsic to these insects were shown to play a key role. Insect immune response, for example, impacts sand fly vector competence to Leishmania. The study of such parameters has been limited by the lack of methods of gene expression modification adapted for use in these non-model organisms. Gene downregulation by small interfering RNA (siRNA) is possible, but in addition to being technically challenging, the silencing leads to only a partial loss of function, which cannot be transmitted from generation to generation. Targeted mutagenesis by CRISPR/Cas9 technology was recently adapted to the Phlebotomus papatasi sand fly. This technique leads to the generation of transmissible mutations in a specifically chosen locus, allowing to study the genes of interest. The CRISPR/Cas9 system relies on the induction of targeted double-strand DNA breaks, later repaired by either Non-Homologous End Joining (NHEJ) or by Homology Driven Repair (HDR). NHEJ consists of a simple closure of the break and frequently leads to small insertion/deletion events. In contrast, HDR uses the presence of a donor DNA molecule sharing homology with the target DNA as a template for repair. Here, we present a sand fly embryo microinjection method for targeted mutagenesis by CRISPR/Cas9 using NHEJ, which is the only genome modification technique adapted to sand fly vectors to date.

Sand Fly Phlebotomus papatasi Embryo Microinjection for CRISPR/Cas9 Mutagenesis

This protocol details the steps of CRISPR/Cas9 targeted mutagenesis in sand flies: embryo collection, injection, insect rearing, and identification as well as selection of mutations of interest.

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to ...