Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

1. Regular maintenance of Harpegnathos saltator colonies

1. Maintain wild-type colonies of H. saltator in transparent plastic boxes in an ant rearing room at 22-25 °C and a photoperiod of 12 hours light: 12 hours dark (12L:12D) lighting schedule.
   1. Use small boxes (9.5 x 9.5 cm²) to rear individual workers or small colonies. Use medium boxes (19 x 13.5 cm²) or large boxes (27 x 19 cm²) to rear larger colonies (Figure 1).
   2. To create nest boxes, use plaster to make floors for the boxes. As the wet plaster is drying in the medium and the large boxes, press a foam block into the plaster a few centimeters deep and a few centimeters from the back of the box to designate a lower nest region. Once the plaster has dried, cover the designated nest region with a square piece of glass.
      NOTE: In small boxes, there is no need to designate a lower nest region. If ants are neglecting to use the designated lower nest region, it may help to cover the glass with a square piece of red cellophane. This gives the impression of a dark underground space resembling nests that H. saltator use in the wild and may encourage them to move their brood to the designated region.
2. Feed colonies with live crickets twice per week.
   NOTE: Colonies should be fed enough that they consume all crickets prior to their next feeding. Feed any isolated ants and mutant ant colonies with crickets pre-stung by workers of a regular colony 2-3 times every week.
3. Apply water regularly to the plaster nest box flooring using a wash bottle.
   NOTE: The plaster should be moist enough that it does not feel dusty to the touch, but it should be dry enough that all added water is absorbed by the plaster. It is important that the nests are not watered excessively. On average, nest boxes will need a small amount of water added once per week.
4. Whenever feeding occurs, remove trash and dead individuals. Freeze all waste and dead ants overnight at -30 °C before disposing of these materials as regular garbage.
5. Add a pinch of dried sawdust to colonies periodically; this helps larvae as they undergo pupation and helps workers keep the nest box clean.

2. Preparation of quartz glass microinjection needles

1. Use a micropipette puller to pull glass microinjection needles.
2. Select the glass to be pulled. Ensure that the glass being used has been stored in a dust-free and clean environment.
   NOTE: Here, thin-walled filamentous quartz glass with an outer diameter of 1.0 mm, inner diameter of 0.5 mm, and length of 7.5 cm has been used to produce microinjection needles. If injecting a soft-bodied embryo, borosilicate needles may also be applicable, but borosilicate needles are not able to penetrate hard chorion.
3. Set the parameter settings of the puller. Use a two-step process to pull microinjection needles for H. saltator embryos: parameters for the first step include heat of 575, filament of 3, velocity of 35, delay of 145, and a pull of 75; parameters of the second step include heat of 425, filament of 0, velocity of 15, delay of 128, and a pull of 200. Following the second step, ensure that the resulting needle has a 2 mm taper and a tip of 0.5 µm (Figure 2).
   NOTE: This set of parameters, along with parameters for other needle types, can be found in the Operation Manual¹⁷. Manuals for pipette pullers often provide recommended parameters for a variety of techniques. Some trial and error may be required to determine which parameters generate the best needles for specific needs. A short taper is ideal for H. saltator injections, as it can penetrate the hard chorion of H. saltator embryos. If injecting a soft embryo, such as one that has been dechorionated (e.g., Drosophila), a longer taper around 10 mm may yield better results. Operation manuals for pipette pullers usually provide specific parameters for pulling needles suitable for different needs. It is important that gloves are worn while handling glass filaments and pulling needles. Oils from bare hands may transfer to the glass if gloves are not worn.
4. Once parameters are set, use a micropipette puller to pull needles for microinjection. Ensure that needles are kept in a dust-free and clean environment until used.
   ​NOTE: It is recommended to use freshly pulled microinjection needles. If pre-pulled needles are used, they should be stored properly in a box to prevent damage of needle tips and potential contamination. Needles that have undergone long-term storage are not recommended for embryo injections.

3. Preparation of microinjector

1. Use a microinjector to inject desired materials into ant embryos.
2. Prepare the microinjection mixture of Cas9 proteins and in vitro synthesized small guide RNAs (sgRNAs)¹⁰. Keep the mixture on ice until it is time to load a microinjection needle. When not in use, store the microinjection mixture at -80 °C.
   NOTE: The concentrations vary in different species. High concentration may cause high mortality, whereas low concentration may reduce efficiency. We use 0.2 µg/µL Cas9 proteins and 0.2 µg/µL sgRNAs for H. saltator embryo injection. Design of our sgRNAs followed a previously established protocol¹⁸. Gene sequences were obtained from DNA databases. The genome sequence of H. saltator was also previously reported^19,20.
3. Adjust the injection parameters for H. saltator embryos: an injection pressure of 140 hectopascal (hPa), a constant pressure of 70 hPa, and a time of 0.4 seconds. Adjust constant pressure such that material only flows in one direction. Adjust the injection pressure and time only if no material is flowing from the needle into the embryo.
   NOTE: If injecting a different type of embryo, the primary parameter that may change is constant pressure, which is responsible for ensuring that fluid from the embryo does not flow back into the needle. Volume is not controlled for in this protocol. Setting the parameters of the microinjector is sufficient for obtaining consistent injections.
4. Load a microinjection needle with 2 µL of the mixture using microloader pipette tips. Do this slowly to ensure that no bubbles are formed in the mixture.
   NOTE: If bubbles form, it may be difficult to maintain consistent microinjections.
5. Break just the tip of the needle by breaking along the edge of the tape such that a narrow taper is still maintained. Ensure that the needle is broken just enough that the tip is opened, but not so much that the taper is broken off.
   NOTE: Importantly, if the opening of the needle is too wide after breaking, the user will see liquid run out of the needle when mounted to the pressurized microinjector prior to application of injection pressure. Some microinjection protocols advise breaking needles by cutting the tip with scissors. This method is not advised, as scissors may cause the tip of the needle to shatter. Injecting embryos with a shattered needle will be highly damaging.
6. Mount the needle to the micromanipulator

4. Injection of embryos

1. Select embryos for microinjection from the syncytial stage: the time during development in which nuclei divide without cytokinesis.
   NOTE: This is the ideal time during development for genome editing by microinjection, as discovered previously in Drosophila²¹. H. saltator embryos pass the syncytial stage and reach cellularization around 36 h after egg deposition¹⁰. Higher efficiency is achieved if younger embryos are used for injections.
2. Line embryos on a piece of double-sided tape stuck to a glass microscope slide. Ensure that embryos are secured well to the tape to prevent movement during injection. Lay the embryos in a vertical orientation, such that the lateral side of the embryo is at the edge of the tape (Figure 3). Place the slide and lined embryos onto the stage of the microscope at a designated microinjection workstation.
   NOTE: It is recommended that the embryos are arranged such that successive injections can be performed by moving the slide on the stage instead of adjusting the needle position with every injection. This allows for successive injections to be performed more efficiently.
3. Align the needle with the first embryo to be injected using the micromanipulator (Figure 4a).
4. Laterally puncture the needle into the first embryo along its dorsal/ventral axis under a microscope.
5. Inject the microinjection mixture. Look for slight movement of the embryo, indicating an increase in internal pressure due to the injected liquid. Additionally, watch for the formation of a small droplet containing visible trace of tissue and/or lipid on the outer membrane of the embryo (Figure 4b).
   ​NOTE: Presence of trace of tissue and/or lipid in the droplet indicates that the needle has successfully punctured both chorion and vitelline membrane of the embryo. If traces of these materials are not present, the injection was not performed successfully and should be repeated. After a few seconds, the droplet will be reabsorbed by the embryo and will no longer be visible.
6. Gently remove the needle from the embryo immediately, and proceed to the next by adjusting the position of the microscope slide. Repeat until all the embryos have been injected.
7. Once all embryos on the slide have been successfully injected, transfer the slide to a humid box for 1 hour to give the embryos time to recover from the injection process prior to being removed from the slide.

5. Rearing of injected embryos

1. After 1 hour of incubation in a humid box, gently remove the injected embryos from the tape using featherweight forceps, and transfer them to a tube filled with a small amount of 70% ethanol. Invert the tube several times to transfer the embryos to the bottom of the tube. Repeat the ethanol wash once, followed by three washes with autoclaved water.
2. Using a small and soft paintbrush, transfer all injected embryos to 1% agar plates with 2% Antibiotic-Antimycotic. Apply the Antibiotic-Antimycotic after poured agar plates have cooled by spreading over the surface of the plate using a cell spreader. Seal the agar plate with parafilm to prevent agar desiccation.
   NOTE: Do not attempt to return injected embryos to a colony after injections as workers may destroy most injected embryos. Survival is therefore optimized by allowing embryos to develop on agar plates outside of a normal colony environment.
3. Incubate the agar plates at 25 °C for approximately 4 weeks. Check regularly for hatching.
4. Once the first embryo has hatched into a larva, return all embryos and larvae to a nest box with a few young nurse workers to care for the hatchlings. Feed using crickets pre-stung by a larger wild-type colony, remove waste products, and add water following the same protocol discussed in Section 1.
   NOTE: Small boxes (9.5 x 9.5 cm²) are ideal for such colonies. H. saltator reproduces well in captivity. Therefore, mutant embryos can be reared to adulthood. Isolation of mutant adult(s) induces transition to the reproductive gamergate stage. Controlled crosses are used to establish mutant colonies with heterozygous or homozygous individuals (Figure 5).