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.

To date, CRISPR has only been used in a handful of Hemipterans. The protocol outlined here could help researchers apply CRISPR and germline transformation in other Hemipteran species. This protocol was developed specifically for insect embryos that are sensitive to desiccation.

Many steps in the protocol could potentially improve outcomes in other insect species. The ability to generate transgenic planthoppers opens the door for novel control methods, such as creating insects that can no longer transmit viruses. This technique will be revolutionary for studying gene function in other pests, such as the western flower thrips.

We have adapted this CRISPR technique for thrips embryos with promising results. We find that practice in this system is essential for success. The methods shown here, when performed routinely, should allow for flexible use in a variety of settings.

To make a chamber to hold the adult Peregrinus maidis insects, cut a hole in the bottom of a one-ounce cup and glue a screen over the hole for air exchange. For the selection of one-week-old adult female insects, aspirate the colony into a 15 milliliter conical tube for one hour, before briefly chilling the insects on ice. Identify the females by the presence of the ovipositor, which is typically darker than the rest of the abdomen, on the ventral side of the abdomen.

Transfer the females into the cup as they are collected. When 15 females have been selected, seal the cup with a five by five centimeter piece of paraffin wax film. Apply 400 microliters of a 10%sucrose solution to the top of the film, and place a second five by five centimeter piece of film over the solution.

Place the adult chamber on an egg collection dish with the plastic paraffin wax film side in direct contact with the oviposition medium, and wrap the entire egg-laying chamber with plastic wrap without covering the air holes. Then, place the chamber at 25 degrees Celsius with 70%humidity and 14 hour light, 10 hour dark cycle. For embryo collection, after the desired egg lay period, apply a one by 15 millimeter strip of double-sided tape onto a 22 by 30 millimeter cover slip and place the cover slip onto the oviposition medium of the egg-laying chamber.

Using a fine brush and a dissecting microscope, move individual semi-transparent embryos from the agar surface to the double-sided tape. When about 25 embryos have been selected, arrange the banana-shaped embryos on their sides with the larger ends stuck on the tape. For microinjection of the embryos, first, place the cover slip onto a 100 by 15 milliliter Petri dish flush with 1%agar under a dissecting microscope inside a humidified hood.

To check the injection pressure, place the tip of a quartz injection needle in a drop of water and initiate the injection cycle. After confirming the pressure setting, approaching from the left side of the cover slip, insert the needle into the larger end of one embryo. Deliver the injection solution into the egg and quickly pull out the needle.

When all of the eggs have been injected, place the cover slip onto the surface of a new 1%agar dish and place the dish into a humidity chamber. After six days, use clean water and fine brush to transfer any surviving embryos to a 35 by 10 millimeter Petri dish containing water-moistened filter paper. Seal the Petri dish with plastic paraffin wax film and place the hatching chamber in a 25 degree Celsius incubator.

After six to eight days, use a fine brush to transfer any emergent first instar nymphs to a Petri dish of leaf clippings and seal the dish with plastic paraffin wax film. After 48 hours at 25 degrees Celsius, use a fine brush to transfer all of the two-day-old nymphs to a rearing cage with corn plants. If injectees with visible phenotype are recovered in sufficient numbers, cage these insects separately to maximize the recovery of the target trait in the next generation.

Rear the insects under the appropriate temperature and humidity conditions with regular transfers to fresh corn plants as necessary, and screen the progeny regularly for expected phenotypes, placing individuals exhibiting the desired phenotype into their own cages to establish homozygous lines. In this representative analysis, a total of 6, 483 eggs were collected from a total of 645 females in four weeks. Females typically start laying eggs after two days and provide most of the eggs from days four to six, with the oviposition activity slowing down by day nine.

Cas9 nine injection does not affect P.maidis development, as the developmental rates for embryos treated with injection buffer, injection buffer supplemented with Cas9, and injection buffer supplemented with Cas9 and three guide RNAs, is similar. The hatch rates for the buffer and Cas9 controls are also similar. However, the hatch rates of individuals receiving the three-guide mix is typically relatively lower.

In this representative experiment, of the 71 guide-injected individuals that developed, 23 showed some degree of pigment loss and nine hatched, resulting in a knockout rate of about 30%The three-guide mix is expected to remove approximately 180 base pairs from the white locus, as observed in the PCR products amplified from genomic DNA isolated from injected individuals, as well as in the associated sequence data generated from those products. We recommend the use of beveled needles to minimize injection trauma. If the embryos appear to leak following injection, examine your needle tips for better results.

Continued practice will increase embryo survival. To date, there are no publications describing germline transformation in Hemipterans. However, we have used this protocol to make Cas9-expressing planthoppers.

Overall, the ability to perform germline transformation in Hemipterans opens the door to a wide range of genetic pest management strategies, thereby reducing the use of insecticides.