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

Summary

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.

Abstract

The corn planthopper, Peregrinus maidis, is 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.

Introduction

The corn planthopper, Peregrinus maidis, is an economically important pest of maize^1,2,3. They cause direct physical damage to the plant, both while feeding with their piercing-sucking mouthparts, and during reproduction when they lay their embryos directly into plant tissue^2,4. Despite the multiple routes of direct damage to crops, the greatest impact these insects have on crop health is indirect, by acting as the vector of maize mosaic virus (MMV) and maize stripe virus^5,6. MMV is capable of replicating in the body of its P. maidis vector, allowing the virus to persist in individual insects through the entirety of their lives, so they can continue to spread the virus to new host plants^7,8. The most common methods for controlling P. maidis, and thus the viruses it vectors, are insecticides.

Unfortunately, mismanagement of these products has caused development of resistance in the target pest as well as pollution of the environment⁹. Therefore, new strategies are needed to reduce crop losses from this insect/virus-pest combination. Previous work demonstrated that RNA interference (RNAi) could be an effective control method for P. maidis because they are susceptible to downregulation in gene expression even when ingesting double-stranded RNA (dsRNA)¹⁰. However, the most effective way to administer dsRNA in the field would be through the plants the insects feed on; hence, crops could still be susceptible to any viruses the insects are already carrying. With the advent of CRISPR/Cas9 genome editing, new pest control strategies are possible, including Cas9-based gene drive^11,12, which could be used to reduce the size of a pest population, or to replace said population with individuals resistant to the viruses they vector.

However, development and deployment of any type of gene-drive system will require the development of transgenic techniques. Such methods were not necessary for carrying out RNAi experiments in P. maidis because dsRNAs and/or siRNAs are presumed to be able to cross cell membranes due to the efficiency of RNAi in P. maidis^10,13. This is not true for the DNAs and/or proteins employed in traditional transgenesis or in Cas9-based gene editing, either of which would be a precursor to creating insects carrying a gene drive. To accomplish gene editing or other forms of germline transformation, these DNAs and proteins are ideally microinjected into embryos during the syncytial blastoderm stage, prior to when the insect embryo cellularizes. Timing is critical, because the syncytial stage is the earliest part of development^14,15. As P. maidis females preferentially lay their eggs in plant tissue, extracting sufficient quantities of precellular embryos for microinjections can be labor-intensive and time-consuming. Therefore, new techniques were developed to quickly collect and microinject P. maidis embryos prior to cellularization.

Protocol

1. Colony-level rearing of P. maidis adults

1. Plant a minimum of four pots of corn per week per rearing cage, with 3-4 seeds per pot. Grow in an insect-free environment.
2. When plants are ~5 weeks old, place inside a 30 cm x 30 cm x 60 cm cage.
3. Obtain a sufficient quantity of P. maidis adults (~500) from a research laboratory or the wild, and place into an insect-proof cage with 9-12 corn plants (3-4 pots).
4. Maintain the colony in an insect-rearing incubator at 25 °C (± 1 °C), with at least 70% humidity and a 14:10 light cycle.
5. To generate an age-calibrated colony, remove all initial adults after four days of egg laying, and allow the embryos laid in the cage to hatch and age naturally.
6. Move 5-week-old P. maidis insects (adults) to fresh corn plants for weekly subculture by collecting with an aspirator (Figure 1). Then, release the adults into a clean cage with fresh corn plants. To maintain a steady supply of young adults for experimental purposes, prepare fresh age-calibrated cages every week.
7. Water the pots in the cages twice daily. Periodically clip stalks, remove decaying plant material, and replace with fresh corn pots as needed.
   ​NOTE: With proper maintenance, a colony can last ~5 weeks (i.e., long enough for embryos laid in the cage to become adults).

2. Agarose-based egg-laying chamber

1. Make egg collection dishes (oviposition medium) by pouring 1% w/v agarose in water into clean 100 mm x 15 mm Petri dishes. Store the oviposition medium at 4 °C after it solidifies.
2. Prepare 10% w/v sucrose solution for feeding the adults. Store the sucrose solution at -20 °C for up to a month.
3. Make a chamber to hold the adults by cutting a hole in the bottom of a 1 oz cup (see the Table of Materials) and gluing a screen over the hole for air exchange (Figure 2).
4. Cut plastic paraffin wax film into 5 cm x 5 cm squares; set aside 2 squares for each cup.
5. Collect ~15 1-week-old adult females from an age-calibrated P. maidis colony. To select females, examine the ventral side of the abdomen, and look for the ovipositor, which is typically darker than the rest of the abdomen (Figure 3). Hold adults for up to one hour in a 15 mL conical vial if setting up multiple egg-laying chambers. Chill the insects briefly on ice prior to sexing and transfer to the adult container.
   NOTE: This examination can be done without a microscope. Adult females that have had time to feed and mate also typically have larger abdomens than adult males and are more docile; hence, they can more easily be selected from a cage population.
6. Transfer the females into an adult container, and seal the cup with 1 layer of plastic paraffin wax film by evenly stretching it 3-4 times its original size (Figure 4A,B).
7. Apply 400 µL of 10% w/v sucrose solution to the top of the plastic paraffin wax film seal, and add a second layer of plastic paraffin wax film, stretching the plastic paraffin wax film exactly as above (Figure 4C,D).
   ​NOTE: The sandwich of stretched plastic paraffin wax film pressurizes the sucrose solution, which is very important for adult feeding, but will not prevent females from piercing their ovipositors all the way through into the oviposition medium.
8. Place the adult chamber on an egg collection dish with the plastic paraffin wax film side directly on the oviposition medium, and wrap the entire egg-laying chamber with plastic wrap without covering the air holes, as these are required for air exchange (Figure 5).
9. Incubate each egg-laying chamber at 25 °C with 70% humidity and a 14:10 light cycle.
10. Change the sandwich of plastic paraffin wax film and 10% w/v sucrose solution daily, and remove any water that accumulates inside the cup.

3. Embryo collection and alignment in a high-humidity environment

1. Set up a stereomicroscope-based microinjection system in a humidified space or hood (humidified hood; Figure 6) to ensure the working environment achieves at least 70% humidity throughout the microinjection process.
2. Check the oviposition medium for eggs after the desired egg-lay period. Do this in a humidified hood or another humid environment.
   NOTE: The egg-lay period typically used was overnight, from 6 PM to 10 AM, lasting ~16 h.
3. If any eggs are laid in the agarose, use fine forceps to carefully dig them out, and place them on the surface of the agarose to keep them moist (Figure 7A).
4. Apply a strip of 1 mm x 15 mm double-sided tape on a 22 mm x 30 mm coverslip (Figure 7B). Place the coverslip tape-side up on the oviposition medium (Figure 7C).
5. Pick up each individual egg from the agar surface, and move to the double-sided tape using a fine brush. Remove any eggs that are completely white or have black coloration on them. Healthy eggs will be the semi-transparent.
6. Place the banana-shaped eggs on their side, with the larger end stuck on the double-sided tape (Figure 7D).
   ​NOTE: Always keep the eggs in a high-humidity environment, such as a Petri dish cast with a layer of 1% agar on the bottom.

4. Preparation of CRISPR reagents and injection needles

1. Pull quartz needles using a Flaming/Brown type micropipette puller.
2. Bevel the quartz needles using a micropipette beveler.
3. Use double-sided sticky tape to secure pulled needles in a clear container, such as a Petri dish, until ready to use.
4. Prepare the injection solution by combining 0.5 µL of Cas9 protein (5 µg/µL stock solution) and 0.5 µL of sgRNA (4 µg/µL stock solution; see the Table of Materials) with 1 µL of phenol red buffer in a final volume of 5 µL. To precipitate particles that could clog the needle, vortex the solution briefly, and centrifuge for 3 min at maximum speed.
5. Backfill the injection needle, taking care to leave the injection mix near the tapered end of the needle. Remove bubbles, if any, from the tip of the needle.
6. Carefully place the backfilled needle into the needle holder, and tighten the stainless steel collar to hold the needle securely in place during microinjection.
7. Generate a reliable flow of injection solution from the needle by gently stroking the beveled tip with a fine, dampened paintbrush, while delivering bursts of air pressure to the needle with the injection system.
   ​NOTE: The needle is ready for injection when the injection mix can leave the tip in small amounts.

5. Microinjection and post-injection care

1. Prepare a microinjection platform by filling a clean 100 mm x 15 mm Petri dish with 1% agar to form a level layer of agar that is flush with the top of the dish.
2. Place a previously prepared coverslip with ~25 embryos upon the agar platform (Figure 8A).
   NOTE: All injection steps must be performed inside a humidified hood (~70% humidity).
3. Check the injection pressure by placing the needle tip in a drop of water and initiating the injection cycle.
   NOTE: A small amount of injection solution should disperse into the water if the pressure setting is correct (Figure 8B).
4. Insert the needle into the larger end of the embryo, approaching from the left side of the coverslip (Figure 8C). Deliver the injection solution into the egg, and pull out the needle quickly.
5. After all eggs are injected, place the coverslip on the surface of a new 1% agar dish, and transfer the dish to a humidity chamber (Figure 9).

6. Incubating and hatching of embryos

1. Put the hatching chamber in a 25 °C incubator for 6 days.
2. Transfer any surviving embryos, using clean water and a fine brush, to a 35 mm x 10 mm Petri dish with water-moistened filter paper covering the bottom of the dish. Seal the Petri dish with plastic paraffin wax film, and hold at 25 °C to allow the embryos to hatch. Start checking the embryos 6 days post-injection for survival.
   NOTE: First instar nymphs will start hatching around day 8.
3. Transfer nymphs, using a fine brush, to a Petri dish containing leaf clippings. Cover the dish, and seal with plastic paraffin wax film.
4. Incubate the sealed dish of hatchlings on leaf cuttings for 48 h at 25 °C.
5. Transfer all 2-day-old nymphs from a round of injections to a rearing cage with corn plants using a fine brush. If injectees with visible phenotype are recovered in sufficient numbers, rear them separately to maximize the recovery of the target trait in the next generation. Otherwise, perform mass-mating of all the injectees.
   NOTE: Place hatchlings gently in the whorl of the corn plant to provide refuge and ensure proper humidity of their immediate environment.
6. Rear the insects in the conditions described above, ensuring proper temperature, humidity, and regular transfers to fresh corn plants.
7. Screen progeny for expected phenotypes. Place individuals exhibiting the desired phenotype into their own cage to establish homozygous lines.

Discussion

Egg-lay quality and nutrition
Recently, researchers working with a related species, Nilaparvata lugens, obtained the eggs they used for microinjections directly from the leaf, keeping the injected eggs in the leaf tissue until they hatched¹⁷. While this leaf-based method provided a more natural environment for embryonic development, it also increased the chances for infections and for egg damage during the removal process. The artificial oviposition system presented here provides a more uniform environment and reduces the chances of damage to the eggs from handling. By setting up the oviposition cups on Friday, the majority of the oviposited eggs were collected during a typical work week, to the benefit of those doing the microinjection work. One caveat to this method, however, is that the lack of nutrients in the 10% sucrose solution diet will eventually affect the health of the insects, and females in the cups usually start to die off after only 10 days. Egg quality also begins to drop off after 6 days, as evidenced by an increase in dead or unhealthy looking eggs. As a result, it is important to be selective of the eggs used for microinjections and to not keep the females after day 6.

Survival rate and humidity
Two factors seem to be critical for embryonic survival through the microinjection process. The most challenging aspect of handling P. maidis embryos is keeping them from desiccating after removal from the oviposition medium and throughout microinjection. As the eggs are typically laid inside plant tissue, they lack an adequate shell to prevent dehydration. Even in the humidified hood whole sets of eggs were lost due to desiccation. However, excessively high humidity could also affect the microinjections if water accumulated on the double-sided tape or on the scope. Unfortunately, egg dehydration was usually not easy to notice during the microinjection process, and they frequently appeared normal until 2 or 3 days later, when they turned completely transparent, showing no signs of development.

Needle quality also appears to play an important role in survival. The needle should be beveled to minimize unnecessary damage to the egg. When the needle is blocked, using the clearing function on the injector while gently stroking the tip of the needle with a dampened paintbrush (see step 4.7) typically returned the needle to a functional state. Regardless, putting only tiny amounts of injection solution (~0.25 µL) into each needle and switching to a new needle every few slides (~50-60 eggs) is recommended to ensure needle quality is maintained throughout the injection process.

Successful generation of a knockout phenotype
To successfully transform the germ cells, embryo microinjections usually have to be done as early as possible before cellularization. Depending on the insect species, the time window for completing the microinjections ranges from only a couple of hours to as long as a full day^14,15,20. It is still unclear when P. maidis embryos undergo cellularization. Cas9-mediated knockout was tested on embryos as young as 4 h post-egg-lay (pel) to as many as 16 h pel, and the expected phenotypes were observed in all experiments, suggesting that all microinjections were performed within the precellurization window.

The P. maidis ortholog of the eye-color gene, white, was selected because the knockout phenotype was expected to be easy to screen in injectees due to its cell-autonomous nature. Indeed, as expected, both mosaic and total knockouts were clearly identifiable among embryos receiving the injection mixture containing Cas9 and guide RNAs. Unfortunately, no injectees with complete knockout hatched, and a mass mating of surviving injectees failed to generate white-eyed progeny. However, a mutant line was later successfully generated by targeting a different gene (Klobasa et al., in progress). This would suggest that the failure to establish a white mutant line is most likely due to either off-target effects (i.e., Cas9 cutting important regions elsewhere in the genome) generating a closely-linked lethal mutation, or to an unpredicted critical role for white in P. maidis.

Phenotypic and molecular data (Figure 8 and Figure 9) affirm that a significant knockout in the white locus was created in a sample of injected embryos, which would result in total loss of gene function. Moreover, while mutations in white are viable in some species, there is precedent for reduced white activity being detrimental^21,22. That said, off-target effects cannot be completely ruled out. Predicting likely off-targets requires accurate genome sequence data²³, which the current state of genomic resources in P. maidis makes impossible to do at this time. Regardless, with these new methods, testing other target genes can be done confidently, even moving towards more traditional transgenesis in an effort to bring new genetic tools to this pernicious pest.

Materials

Name	Company	Catalog Number	Comments
1 oz Containers	Dart	P100N	Adult container for egg-laying setup
15 mL Conical Tubes	Olympus	Genesee 28-103	Serves as collection tube on vacuum aspirator setup
15 mL Conical Tubes	Olympus	Genesee 28-106	For making 10% sucorose solution and for holding adults when chilling before screening
Aspirator	Bioquip	1135A	For handling planthoppers
Vacuum Aspirator	Fischer Technical	LAV-3	Vacuum for aspirating larger numbers of insects
Blue Spectrum LED Lights	Home Depot	GLP24FS/19W/LED	Grow lights for potted corn plants hoppers are feeding on
Cas9	TrueCut Cas9 Protein v2	A36498	Endonuclease for cutting planthopper genes
Clear Vinyl Tubing	Home Depot	3/8 in. I.D. x 1/2 in. O.D. x 10 ft.	Connects collection tube to pump on vacuum aspirator setup
Corn planthoppers	North Carolina State University	N/A	Request from Dr. Anna Whitfield's lab
Cotton balls	Genessee	51-101	Serves as a filter/insect catcher in collection tube on vacuum aspirator setup
Double sided tape	Scotch Double Sided Tape	NA	Holding eggs for microinjection
Early Sunglow corn	Park Seed Company	05093-PK-N	Corn for rearing planthoppers
epTIPS Microloader Tips	Eppendorf	C2554691	Backfilling needle loading tips
Femtojet Microinjection System	Eppendorf	5247	Controls injection pressure (12-20 psi, depending on needle bore size)
Nutri-Fly Drosophila Agar	Genessee	66-103	Substrate for everything except egg-laying dish
Fine forceps	Bioquip	4731	Egg handling
General Purpose LE Agarose	Apex	20-102	Substrate inn egg-laying dish (oviposition medium)
Guide RNA 1 - GGUUCAUCGCAAAAUAGCAG	Synthego	CRISPRevolution sgRNA EZ Kit (1.5 nmol)	RNA guides for targeting planthopper white gene
Guide RNA 2 - UCUGAAAUCACUGGCCAAUA	Synthego	CRISPRevolution sgRNA EZ Kit (1.5 nmol)	RNA guides for targeting planthopper white gene
Guide RNA 3 - GAGGGCAGAGUCGCUUUCUU	Synthego	CRISPRevolution sgRNA EZ Kit (1.5 nmol)	RNA guides for targeting planthopper white gene
Humidifyer	Homedics	UHE-CM45	For providing humidity in humidified hood
Humidity chamber	Billups-Rothenberg	MIC-101	For holding injected embryos until hatching
Insect rearing cages	Bioquip (special order)	Close to 1450 L (has plastic front and mesh fabric sides)	Cage for planthoppers on corn
Laser-based Micropipiette Puller	Sutter Instruments	P-2000/G	For making injection needles / Heat = 700, FIL = 4, VEL = 40, DEL = 170, PUL = 160
Leica M165 FC Fluorescence Stereomicroscope	Leica	M165 FC	Planthopper screening
Microinjection Scope	Leica	MZ12-5	Microinjection scope outfited with an XY stage
Micromanipulator	Narishige	MN-151	For positioning microinjection needle
Micropipette beveler	Sutter Instruments	FG-BV10-D	For beveling injection needles / Used 'fine' graded plate at 20° angle
Microscope Stage	AmScope	GT100 X-Y Gliding Table	For positioning and moving embryos under microscope
Miniature Paint Brush	Testor #2 8733	Sold in 3 pack 281206	Fine paintbrushes for embryo handling
Needle Holder	Narishige	HI-7	For holding the microinjection needle
Percival Incubator	Percival	I41VLH3C8	Rearing injectees until hatch
Petri Dishes (100 x 15 mm)	VWR	89038-968	Making agar dish for egg-lay
pGEM-T Easy Vector System I cloning kit	Promega	A1360	Cloning Pm white target site
Phenol Red	Sigma	143-78-8	Microinjection buffer
Plain Microscope Slides or coverslip	Fisher Scientific	12-549-3	Hold eggs for microinjection
Plasmid DNA Midi Kit	Zymo	D4200	Purification of injection-ready plasmid DNAs
Plastic paraffin film	Pechiney Plastic Packaging	PM-996	Roll size 4 in. x 125 ft
Plastic wrap	Glad ClingWrap Plastic Wrap	NA	Wrap the entire egg-laying chamber
Primer - PmW CRISPR check F1 - AAGGAATTTCTGGAGGTGAAA	IDT	25 nmole DNA Oligo	First-round Primer for amplifing across target site within the Pm white gene
Primer - PmW CRISPR check R1 - GATTCCTCGCTGTTGGGT	IDT	25 nmole DNA Oligo	First-round Primer for amplifing across target site within the Pm white gene
Primer - PmW CRISPR check F3 - TCACAGACCCTGGTGCTAATC	IDT	25 nmole DNA Oligo	Second-round Primer for amplifing across target site within the Pm white gene
Primer - PmW CRISPR check R3 - GTCCACAATCCACACTTCTGA	IDT	25 nmole DNA Oligo	Second-round Primer for amplifing across target site within the Pm white gene
Quartz capillaries	Sutter Instruments	QF100-50-10	For making microinjection needles / O.D. 1 mm, I.D. 0.7 mm, 10 cm length
Screen (White Organza Fabric)	Joann Fabrics	16023889	For covering the adult container
Sparkleen	Fisher Scientific	04-320-4	Wash dishes
Sucrose	Fisher Scientific	BP220-1	To make 10% sucorose solution