Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

1. N. Vitripennis Colony Rearing

1. 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.
   1. Maintain the wasps at 25 ± 1 °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 °C will slightly lower the developmental timing of the embryos.
2. Allow wasps to mate for at least 4 days prior to injections.
   1. 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.
   2. For the following two days, remove the blowfly pupae to deprive the female wasps of an oviposition site, making them very gravid.

2. Collection and Alignment of N. Vitripennis Pre-blastoderm Stage Embryos

1. Allow female wasps to parasitize (oviposit embryos) into young hosts (S. bullata pupae). To keep hosts fresh, store hosts in 4 °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).
   1. 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 'crater-like' opening (Figure 1B).
   2. 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 (> 1.5 h) should not be injected.
   3. 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.
2. 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.
3. 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.
4. 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 'embryo pick,' which is essentially a half of a pair of ultrafine forceps, can be used instead of a paintbrush.
5. 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'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 previously^21,22. Additionally, do not desiccate embryos, or cover the eggs with halocarbon oil during injection as described previously for N. vitripennis embryo microinjection²².

3. Needle Preparation for Microinjections

1. 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).
2. 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.
3. 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's clay.
4. Bevel the needle tip by slightly touching the tip to the diamond abrasive plate around 10 s at 25 °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.

4. Embryo Microinjection

1. Prepare the injection mixture consisting of genome modification reagents (e.g., transposon + helper transposase, or CRIPSR reagents, etc.), and keep it on ice.
2. Load the injection needle with 2 µL 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.
3. Place the glass slide with the lined embryos onto the stage of a compound microscope.
4. Carefully insert the needle into the posterior end of the embryo with a vertical angle of 25-35 °. Inject 1-5 pL of injection mixture (around 2-10% of the embryo'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.
   1. 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.
   2. 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 (> 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.

5. Transplanting Injected G₀ N. vitripennis Embryos onto Pre-stung Host

1. 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.
   1. 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.
2. Place the host pupae into a Petri dish with damp filter paper and cotton balls, and incubate them at 25 °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.
3. Transfer the host pupae into a new Petri dish at 70% humidity and 25^°C after G₀ embryos hatch.
   1. 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.

6. Screening for Genome Modifications

1. 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.
2. 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.
3. 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.
4. 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.