Genome Editing in the Yellow Fever Mosquito Aedes aegypti using CRISPR-Cas9

Summary

Here, we describe a detailed protocol for genome editing through embryonic microinjection in the mosquito A. aegypti using the CRISPR-Cas9 technology.

Abstract

The emergence of the clustered, regularly interspersed, short palindromic repeats (CRISPR)-Cas9 technology has revolutionized the genetic engineering field and opened the doors for precise genome editing in multiple species, including non-model organisms. In the mosquito Aedes aegypti, loss-of-function mutations and DNA insertions have been accomplished with this technology. Here, we describe a detailed protocol for genome editing through embryonic microinjection in the mosquito A. aegypti using the CRISPR-Cas9 technology, focusing on both the generation of gene knockout and knockin lines. In this protocol, quartz needles are filled with a mixture of guide RNA, recombinant Cas9, and a plasmid containing a DNA cassette encoding a gene for a fluorescent marker, if gene knockin is desired. Embryos at the preblastoderm stage are lined up onto a strip of double-sided sticky tape placed onto a coverslip, which is subsequently mounted onto a glass slide. With the help of a microinjector, the needles are inserted gently into the posterior end of the embryos and a small volume of the CRISPR mixture is dispensed. When the embryos are hatched, the larvae are checked under the fluorescent scope, and the pupae are sex-sorted and separated in different cages. Once the adults emerge, these are reciprocally crossed with wild-type individuals, blood-fed, and placed for egg laying. Once these eggs are hatched, the fluorescent larvae collected represent individuals with stable insertion of the DNA cassette into their genome. These larvae are then grown to the adult stage, outcrossed to wild-type individuals, and then further assessed through molecular techniques to confirm that the exact sequence of the DNA cassette is present at the desired site of the mosquito genome. Homozygous lines can also be obtained by following the provided pipeline of crossing schema and molecular screening of the mutations.

Introduction

Precise genome editing has become arguably easier, but possible, with the establishment of the CRISPR-Cas technologies of molecular scissors¹. These technologies take advantage of a mechanism that the prokaryotic immune system uses to fight against phage infections². Amongst these systems, clustered, regularly interspersed, short palindromic repeats (CRISPR) along with the Cas9 nuclease usually rely upon 20 base pair RNAs, the guide RNAs (gRNAs), with sequences homologous to the targeted DNA, which are followed by an NGG protospacer adjacent motif (PAM) sequence³. The gRNAs loaded onto the Cas9 guide these nucleases precisely to specific target sites in the genome, triggering double-strand DNA breaks³.

DNA double-strand breaks induce the repair mechanisms to patch the double-helix⁴. Whereas any DNA repair is expected to be precise, there exist less accurate DNA repair mechanisms that can leave behind sequence scars and, in turn, loss-of-function mutations⁴. Among the error-prone DNA repair mechanisms, the Non-Homologous End-Joining (NHEJ) can cause frame-shift mutations, including small deletions, insertions, and nucleotide changes (SNPs), that can result in loss-of-function mutation. The Homology Directed Repair (HDR) mechanism, on the other hand, relies upon the homologous chromosome as a template to copy the exact sequence of the undamaged allele and make a perfect repair of the targeted DNA sequence⁴.

Based on this knowledge, the CRISPR-Cas9 technology has been developed to precisely edit genomes, arguably at any sequence containing a PAM site³. In mosquitoes, the CRISPR-Cas9 technology has been used to knock out a variety of genes, through embryonic microinjection of a mixture of Cas9 and gRNAs, taking advantage of the NHEJ repair mechanism^5,6. Similar germline mutagenesis is obtained with the injection of gRNA + Cas9 mix into the hemolymph of adult female mosquitoes⁷. This technology was coined ReMOT control and relies upon a modified version of a Cas9 tagged with a peptide that is taken up by the ovaries through endocytosis during the process of egg development (vitellogenesis)⁷. Knocking in specific gene cassettes into a genome is only possible through embryonic microinjection of a mixture of gRNA and Cas9 (or a plasmid expressing those molecules) along with a plasmid encoding a desired DNA cassette⁸. Taking advantage of the HDR mechanism, the plasmid containing the DNA cassette of interest flanked by homologous sequences (500-1,000bp)^9,10 upstream and downstream of the target site is used as a template to rewrite the double-strand break, copying also the DNA cassette into the target sequence⁹.

The CRISPR-Cas9 technology has been used to knock out multiple genes primarily involved with the sensory systems in the mosquito Aedes aegypti¹¹, but also genes associated with male fertility and female viability (PgSIT) for population control¹². Knocking out target genes has also been accomplished by knocking in genes encoding fluorescent markers into the coding sequences of specific genes^13,14. This strategy has the advantage of not only inducing frame-shift mutations but also allowing the use of fluorescent light to sort the individuals of the new knockout line^13,14. The A. aegypti genome has also been edited with sequences of binary expression systems, such as the Q-system (QF-QUAS)¹¹. Knocking in the gene encoding the QF transactivator downstream to a promoter of a specific gene assures defined spatiotemporal expression of the transactivator^15,16. Once a QF-expressing mosquito line is crossed to another mosquito line containing the binding sites (QUAS) for QF, the latter binds to it and triggers the expression of genes downstream to the QUAS sequence^15,16. This system, overall, allows tissue- and time-specific expression of such effector genes, which can be fluorescent markers used for cell localization or detection of neuronal activity, and even Cas9 nucleases for disrupting genes in specific tissues (i.e., somatic knockout)¹¹.

Given all the information available for A. aegypti genetic transformation, we provide herein a detailed protocol with step-by-step directions to performing genome editing with the CRISPR-Cas9 system through embryonic microinjection. Strategies for generating both knockout, through frame-shift mutations and deletions mediated by NHEJ, and knockin lines, by HDR-mediated gene cassette insertions, are discussed.

Protocol

Details related to the equipment and reagents used in this protocol are listed in the Table of Materials. All animals were handled following the Guide for the Care and Use of Laboratory Animals, as recommended by the National Institutes of Health. The procedures were approved by the UCSD Institutional Animal Care and Use Committee (IACUC, Animal Use Protocol #S17187) and UCSD Biological Use Authorization (BUA #R2401).

1. gRNAs and donor plasmid design

1. For making knockout mutants, design two gRNAs spaced by ~20-100 bp (Figure 1A).
   1. Design 20 bp gRNAs for CRISPR-Cas9, excluding the PAM sequence (NGG) by using an online tool (Figure 1A), such as CHOPCHOP ( https://chopchop.cbu.uib.no/)¹⁷or Benchling (benchling.com) or CRISPOR (http://crispor.gi.ucsc.edu/)¹⁸. Select the most specific and off-target-free gRNAs for experiments, as suggested by the online tool.
   2. Ensure an efficient restriction enzyme cutting site is included within the sequence between the gRNA cut sites (Figure 1A). Include a restriction enzyme site between gRNA cut sites to allow for quick visual confirmation of successful edits.
      NOTE: When the deletion occurs, the restriction site is removed, so the enzyme will not cut the sequence, producing a single, uncut band on a gel to confirm the deletion.
2. For HDR-mediated gene cassette insertions, design multiple gRNAs using the tools mentioned for making knockout mutants and select the most effective one after later evaluation.
3. Design donor plasmids for Homology-Directed Repair (HDR) containing target site homology arms, the cargo DNA sequence, a fluorescent marker, the origin of replication, and an antibiotic resistance gene (Figure 1B).
   1. Choose the homology arms from the upstream and downstream regions of the target site, each spanning 500-1,000 bp¹⁰ (Figure 1B).
   2. Select the cargo sequence, which may include a fluorescent marker, a gene of interest, or a regulatory element (e.g., QF2).
   3. Select a fluorescent marker. Common fluorescent markers used for A. aegypti are included in a DNA cassette containing a promoter, a fluorescent marker-encoding gene, and a 3' UTR sequence. See the discussion for further details.

2. Injection mix preparation

1. Dilute the Cas9 protein with the Cas9 dilution buffer to 1 µg/µL.
   NOTE: Do not thaw and freeze Cas9 more than 2x. It is advisable to make aliquots of the Cas9 protein.
2. Purchase the gRNAs or produce them in-house.
   NOTE: For in-house production, use standard RNA decontamination practices and RNAse-free materials.
   1. Design 4-6 gRNAs and choose the best two gRNAs for injection (see in vitro cleavage assay below).
   2. Design a forward primer that includes the T7 promoter sequence upstream of the gRNA sequence. Use the universal gRNA reverse primer for the non-template PCR reactions, as described below. The overlapping sequence base pairs to the corresponding sequence in the universal reverse primer (bold and underlined), creating a template for the DNA polymerase to amplify their sequences.
      NOTE: For these PCR reactions, the forward primer should contain the T7 promoter (bold), followed by two guanines (important for transcription initiation via T7 RNA polymerase), the 20-nucleotide sequence of the gRNA (N₂₀; without the PAM sequence), and the primer overlapping sequence (underlined).
      Primer forward: 5'- GAAATTAATACGACTCACTATAGGN₂₀ GTTTTAGAGCTAGAAATAGC- 3'
      Primer reverse: 5''-AAAAGCACCGACTCGGTGCCACTTTTT CAAGTTGATAACGGACTAGC
      CTTATT TTAACTT GCTATTTCTAGCTCTAAAAC - 3'
   3. Synthesize the DNA template by non-template PCR. Set up multiple reactions (each containing 12.5 µL of the 2x master mix, 1.25 µL of the 10 µM solution of forward primer, 1.25 µL of the 10 µM solution of reverse primer, and 10 µL of ultrapure water) so that enough PCR product (300 ng) is available for the in vitro transcription reaction. Use the following PCR conditions: initial denaturation for 30 s at 98 °C; 35 cycles of amplification for 10 s at 98 °C, 10 s at 62 °C, and 10 s at 72 °C; final extension at 72 °C for 2 min; and storage at 4 °C.
   4. Confirm amplification of a single DNA fragment (122 base pairs) on an agarose gel (2%).
   5. Clean up the PCR template using a PCR Purification kit, following the manufacturer's recommendations.
   6. Perform an in vitro transcription reaction with a T7 Transcription Kit. Mix 2 µL of 10x reaction buffer, 2 µL each of nucleotides (ATP, CTP, UTP, and GTP), 2 µL of T7 RNA polymerase, 3 µL of the template DNA (100 ng/µL), and 5 µL of ultrapure water. Incubate the mixture at 37 °C for at least 2 h (not more than 16 h) to overnight (12 h).
      NOTE: In this reaction, the T7 RNA polymerase binds to the T7 promoter included in the primer forward sequence, which leads to the transcription of the gRNA. 
   7. Treat the transcription reaction with DNase by adding 1 µL of DNase (mix well) and incubating at 37 °C for 15 min.
   8. Purify the synthesized sgRNAs with a transcription cleanup kit by following the manufacturer's recommendations.
   9. Perform an in vitro cleavage assay to assess the cutting efficiency of the selected gRNAs (Figure 1C).
      1. Design primers to amplify a DNA fragment (500-1,000 bp) flanking the gRNA cutting site.
      2. Set up the PCR reactions as follows: 12.5 µL of the 2x master mix, 1.25 µL of the 10 µM solution of forward primer, 1.25 µL of the 10 µM solution of reverse primer, 9 µL of ultrapure water, and 1 µL of 5 ng/µL template DNA.
      3. Use the following PCR conditions: conditions: initial denaturation for 30 s at 98 °C; 35 cycles of amplification for 10 s at 98 °C, 15-30 s at X °C (ideal annealing temperature for primers), and 35 s at 72 °C; final extension at 72 °C for 2 min; and final long-term storage at 4 °C.
      4. Refer to the manufacturer's guidelines for the best amplification temperatures and time span. Pool at least five reactions or scale up the volumes of the reagents so that enough PCR product (1.5-2 µg) is obtained after PCR cleanup (single band) or agarose gel PCR band purification.
      5. Set up Cas9 cleavage reactions with recombinant Cas9 by incubating the following mixture at 37 °C for 1 h: 1 µL of 10x Cas9 reaction buffer, 0.35 µL of recombinant Cas9 (1 µg/µL), 1 µL of gRNA (100 ng/µL), 6.65 µL of ultrapure water, and 1 µL of 300 ng/µL of template DNA.
      6. Set up a negative control reaction without any gRNA.
      7. Check for cleavage efficiency by running reactions on an agarose gel (1.5-2.0%; Figure 1C).

3. Assembling the donor plasmid

1. PCR-amplify the homology arms from the genomic DNA of A. aegypti.
   1. For primer design, refer to the cloning kit manufacturer's guidelines. Use an online tool (e.g., Benchling), which provides support for plasmid design and assembly using different cloning strategies (e.g., Gibson, Golden Gate, and restriction enzyme-based cloning).
   2. Amplify the cargo sequence from A. aegypti genomic DNA or order a commercially synthesized gene fragment.
   3. Amplify fluorescent markers from plasmids in-house.
2. Ligate DNA fragments from steps 3.1.1 to 3.1.3 into the backbone of an existing plasmid by Gibson assembly. Incubate a mixture of 10 µL of 2x master mix, X µL of all DNA fragments, 10-X µL of ultrapure water at 50 °C for 1 h.
   NOTE: Refer to the manufacturer's guidelines for calculating the ideal ratio of inserts to plasmid backbone.
   1. Calculate the concentrations of each cloning fragment in pico mols: pmols = (weight in ng) × 1,000/(base pairs × 650 daltons).
   2. Use 50-100 ng of plasmid backbone and 2-3-fold molar excess of each insert.
   3. If assembling 2-3 fragments, add 0.02-0.5 pmols of each fragment into the Gibson reaction. If assembling 4-6 fragments, add 0.2-1.0 pmols of each one.
3. Use 3-5 µL of the Gibson assembly reaction to transform E. coli competent cells JM109.
   NOTE: JM109 was chosen for Gibson assembly due to its recA- genotype, which reduces undesirable recombination events and prevents nuclease carryover during cell harvest, ensuring the integrity of the assembled DNA fragments
   ​For plasmids larger than 10 kb, we recommend transformation of DH5α competent cells using the extended protocol, following the manufacturer's recommendations.
4. Expand the transformed bacteria and purify the plasmid using a miniprep kit.
5. For confirmation of correct plasmid assembly, run a diagnostic restriction enzyme digest with the purified plasmid DNA and visualize by agarose gel electrophoresis (Figure 1D,E).
   NOTE: We suggest the use of an online tool for the selection of a couple of restriction enzymes that can cut the plasmid on a single site. Software like Benchling has a built-in tool that performs virtual digestion of plasmid sequences with the selected restriction enzymes, displaying the expected DNA band pattern on a virtual electrophoresis gel (Figure 1D).
   1. Carry out the plasmid restriction enzyme digestion. Incubate a mixture of 1 µL of 10x restriction digest buffer, 0.5 µL of restriction enzyme 1, 0.5 µL of restriction enzyme 2, 1 µL of plasmid DNA (300 ng/mL), and 7 µL of ultrapure water at 37 °C for 2 h.
   2. Run the restriction digest reactions on an agarose gel (1.5%).
      NOTE: The DNA band pattern (Figure 1E) should resemble the virtual digest band pattern (Figure 1D). Plasmids can also be sent for whole plasmid sequencing if service is available.
6. Culture the plasmid clone into 150 mL of LB media.
7. Carry out plasmid maxiprep, following the manufacturer's guidelines.
8. Suspend the plasmid in ultrapure water.

4. Mixing the injection construct

NOTE: The suggested final concentration ranges for each construct are provided in Table 1. Start with a ratio of 2:1:2 (ng Cas9: ng sgRNA: ng donor plasmid) and adjust as necessary to optimize HDR efficiency. Monitoring the outcomes will help identify the most effective combination.

1. For making knockout mutants,
   1. Dilute the aliquot of Cas9 protein to the desired concentration using the Cas9 dilution buffer and dilute the aliquot of gRNA with ultrapure water.
   2. Premix the Cas9 protein with each gRNA to form a Ribonucleoprotein (RNP) complex. Combine the premixed solutions.
2. For HDR-mediated gene cassette insertions,
   1. Dilute and mix Cas9 protein and gRNAs as suggested for making knockout mutants.
   2. Dilute the donor plasmid with ultrapure water.
   3. Combine all the constructs.

5. Pulling and loading microinjection needles

1. Use quartz filament to pull the injection needles (Figure 2). Make sure the filament is 10 cm long, with an outer diameter of 1 mm and an inner diameter of 0.7 mm.
2. Pull the needles using a laser micropipette puller with one of the following two programs:
   Program 1: Heat 805, Filament 4, Velocity 50, Delay 145, Pull 145
   Program 2: Heat 650, Filament 4, Velocity 40, Delay 150, Pull 156
   NOTE: Program 1 results in a thinner but longer needle tip, making it suitable for injections with lower concentration constructs, where a finer tip is beneficial for precision. Program 2 yields a shorter but thicker needle tip, ideal for higher concentration injections, as the construct is thicker and requires a sturdier needle.

6. Embryo harvesting

1. Prepare an electric or manual aspirator to collect mosquitoes and use plastic vials for collection and embryo harvesting.
2. Moisten white circle filter papers and place them on the inside wall or on top of moist cotton in the collector.
3. Place 5-10 female mosquitoes that were blood-fed 5-10 days ago into the collector. Place the collector in the dark for 45 min.
4. Take out the filter papers for embryo harvesting.

7. Embryo line up

1. Use a wildtype strain (Liverpool) of A. aegypti for embryo injection.
2. Select preblastoderm stage embryos, particularly those that are light gray in color, from the harvesting paper (Figure 3).
3. Transfer a few embryos with a wetted brush to the double-sided sticky tape placed on top of a coverslip. Align the embryos in parallel while their surrounds are still wet ensuring they are side by side, with all posterior ends facing the front.
4. Once positioned correctly, allow the environment to dry slightly to secure the embryos in place.
5. Add Halocarbon oil 700 onto the embryos during alignment to prevent desiccation.
   NOTE: Ensure the embryos are not surrounded by water when adding the oil, as this would cause the embryos to float in the oil.

8. Embryo microinjection

NOTE: Injection is conducted at room temperature or at 18 °C. The 18°C temperature is recommended for practical reasons, as it delays embryo development.

1. Set up a microinjector with the following parameters: compensation pressure (Pc) 300 hPa, injection pressure (Pi) 500 hPa. Adjust these conditions whenever necessary.
2. Load 3 µL of the mixed construct into a needle using a microloader.
3. Place the cover glass with aligned embryos on top of a microscope slide (Figure 3). Position the cover glass and slide under the microscope for injection. Make sure the posterior end of the embryo is positioned toward the needle.
4. Place one needle into a needle holder together with a micromanipulator and position the needle at a 10° angle towards the posterior end of the aligned embryos, keeping it stationary (Figure 4A). Gently open the needle under a microscope and lightly touch it with the edge of a coverslip.
   NOTE: Another option is to create a small opening in the needle by tapping it with an embryo. However, a slightly wider needle opening is recommended, as the Cas9 protein's stickiness can clog the needle.
5. Move the slide glass towards the needle for injection (Figure 5).

9. Injected embryo aftercare

1. Use lint-free, disposable wipes to remove the oil surrounding the embryos.
2. Add deionized water to rinse the embryos, and move the embryos onto a wet filter paper. Place the wet filter paper on a wet tissue inside Karat 9 oz cups (Figure 4B and Figure 6A,B).
3. Place wet cotton at the bottom of the cup to maintain moisture (Figure 6B).
4. Preserve the injected embryos on wet filter paper.

10. Embryo hatching and G0 larva screening

1. At least 4 days after injection, transfer the filter paper with embryos to approximately 3 L of deionized water in Sterilite 6-quart pans for hatching.
   NOTE: Eggs typically hatch more efficiently within 2 weeks after injection. Ensure the eggs remain moist during this period. The filter paper should not be too wet, as excessive moisture may lead to early hatching of the embryos. While longer preservation may improve hatching rates, do not exceed a month for optimal results. Using deoxygenated water may also help with hatching.
2. Once the G0 larvae hatch, add fish food mixed with water to the pan as food.
3. Screen the G0 larvae for the fluorescent marker at the 3rd to 4th instar larval stage (Figure 7).
4. Maintain the larvae separately based on their fluorescent status, with fluorescent-positive and fluorescent-negative larvae kept in separate pans.

11. Sex sorting pupae and outcrossing to wild type

1. Separate the injected mosquitoes by sex when they pupate, using size and sex-specific structures at the genital lobe for identification:
   1. Males: Smaller size, a more prominent and pointed genital lobe, and broader paddles (structures at the tail end of the pupae; Figure 8A1,A2).
   2. Females: Larger size, a smaller and less pronounced genital lobe, and narrower paddles (Figure 8B1,B2).
2. Pool fluorescent-positive or fluorescent-negative mosquitoes of each sex.
3. Outcross the pooled mosquitoes with mosquitoes of the opposite sex from the Liverpool strain. Use a 3:1 to 5:1 ratio of wild type to G0 mosquito.
4. Cross the mosquitoes for 4 days.
5. Provide the females with a blood meal after crossing.

12. G1 screening

1. Three days after blood feeding, provide egg cups by placing a paper towel inside the wall of Karat 9 oz cups and adding around 3 oz of deionized water.
2. After 3-4 days, harvest and hatch the G1 embryos. Screen the G1 larvae for the fluorescent marker at the 3rd to 4th instar stages (Figure 7).
3. Collect the fluorescent larvae, suggesting an HDR insertion. When the fluorescent G1 individuals reach the pupal stage, separate them by sex and place each sex in separate cages.
4. Outcross the fluorescent G1 mosquitoes to individuals of the opposite sex of the Liverpool strain.

13. G1 insertion site confirming

1. After providing the blood meal, set up G1 females for single female egg laying by placing a small piece of paper towel in individual Drosophila vials and adding 3 mL of deionized water.
2. For knockout mutants:
   1. After egg laying, water evaporates, and the paper towel dries out, hatch the eggs from females that clearly exhibit knockout mutations and obtain the G2 generation.
   2. Collect the body of the G1 mothers have successfully hatched G2 offspring and extract DNA.
   3. Use the DNA as a template for PCR to amplify the sequence covering the target site. Make sure the primers amplify a DNA fragment of ~200 bp. Set up the reactions (each containing 12.5 µL of the 2x master mix, 1.25 µL of the 10 µM solution of forward primer, 1.25 µL of the 10 µM solution of reverse primer, 9 µL of ultrapure water, and 1 µL of template DNA [5 ng/µL]). Use the following PCR conditions: initial denaturation for 30 s at 98 °C; 35 cycles of amplification for 10 s at 98 °C, 15-30 s at X °C (primers' ideal annealing temperature), and 10 s at 72 °C; final extension at 72 °C for 2 min; and final long-term storage at 4 °C.
   4. Purify PCR fragments and perform restriction enzyme digestion of the PCR fragments by incubating a mixture of 1 µL of 10x restriction digest buffer, 0.5 µL of restriction enzyme 1, 0.5 µL of restriction enzyme 2, 1 µL of PCR fragment (200 ng/mL), and 7 µL of ultrapure water at 37 °C for 30 min.
      NOTE: Certain restriction enzymes can be used in the PCR mix without prior purification.
   5. Visualize the PCR fragment digestion by gel electrophoresis to confirm whether cleavage has occurred.
   6. Sequence the undigested PCR fragment to identify the knockout mutations.
   7. For HDR-mediated insertions, after egg laying:
      1. Collect the body of the G1 mothers and extract DNA.
      2. Use DNA as a template for PCR to amplify the sequence covering the target site.
      3. Perform sequencing of the amplified DNA fragment to confirm the integrity of the inserted DNA cassette.
      4. Hatch the egg and obtain the G2 generation.

14. Expanding the new CRISPR lines

1. For knockout mutant lines:
   1. Set up G2 females for egg-laying, with 20 females per G1. Each G1 represents one line.
   2. After egg laying, confirm knockout mutations in the same manner as for G1.
   3. Proceed with five G1 lines that exhibit clearly identified mutations in both G1 and G2. Hatch the eggs collected from G2 of each line, giving rise to the G3 generation.
   4. From the five outcrossed lines, select two lines with clearly identified mutations and high fitness or the desired phenotype.
   5. Outcross G3 females from each selected line with males from the Liverpool strain.
   6. Set up 50 single females for egg-laying from each line. Confirm knockout mutations as done previously for G1.
      NOTE: Increase the number of females in the setup to maintain diversity.
   7. Repeat the procedures for two additional generations with the two lines (Figure 4J).
2. For new HDR-mediated insertions lines:
   1. Verify the presence of fluorescent individuals in the G2 generation.
   2. Outcross G2 mosquitoes, from each G1 individual, with the Liverpool strain. Each G1 individual represents one line.
   3. Proceed with three more generations of outcrossing. Continuously check the fluorescent marker and observe the fitness¹⁹ of each line (Figure 4x).
   4. Select one line with correct insertions and high fitness and the desired phenotype.

15. Make homozygous lines

1. For knockout mutant lines:
   1. Generation G6:
      1. Hatch the eggs produced by G5 and obtain the G6 generation.
      2. Intercross G6 adults within their respective lines (Figure 4K and Figure 9A).
      3. Set up 50 single G6 females for egg-laying per line, and hatch the eggs as previously described (Figure 9B).
      4. Collect the G6 females that have successfully hatched G7 offspring, extract DNA, and perform PCR and restriction enzyme digestion as done for G1 (Figure 9C).
         NOTE: A simple and inexpensive squishing buffer (SB) can be used at this step for DNA extraction.
         1. Using a handheld grinder, macerate a single mosquito in 100 µL of 1x SB (10 mM Tris-HCl pH 8, 1 mM EDTA, and 25 mM NaCl). Add Proteinase K (200 µg/mL) and incubate at 37 °C for 30 min, followed by 95 °C for 2 min. Spin down at 10,000 × g for 5 min and collect the supernatant²⁰.
      5. Confirm mutation by visualizing the results on a gel.
      6. Discard the eggs produced by females without mutations (Figure 9D).
      7. Maintain larvae from females with confirmed mutations to establish several G7 lineages, and discard broods without mutations (Figure 9D).
   2. Generation G7:
      1. Intercross females and males within each G7 lineage (Figure 9E).
      2. Set up 10 single G7 females for egg-laying per line, and hatch the eggs as previously described (Figure 9F).
      3. Collect the G7 females that have successfully hatched G7 offspring, extract DNA, and perform PCR and restriction enzyme digestion (Figure 9G).
      4. Detect knockout mutations on both alleles or a single allele by visualizing the results on a gel.
      5. Maintain larvae from G7 females confirmed to have knockouts in both alleles, producing the G8 generation (Figure 9H). Discard broods without mutations (Figure 9H).
         NOTE: This step ensures that the females are homozygous. In G8, testing the homozygosity of the progeny confirms whether the fathers are also homozygous."
   3. Generation G8:
      1. Cross G8 adults per each G7 female lineage (Figure 9I).
      2. Randomly pick five G8 males, extract DNA, and perform PCR and restriction enzyme digestion (Figure 9J).
      3. Provide a blood meal and hatch the eggs from G8 females that were crossed with G8 males confirmed to have knockouts in both alleles.
      4. Continue with the G7 lineages where all detected males have knockouts in both alleles, indicating these lineages are homozygous.
2. For HDR-mediated insertions lines:
   1. Hatch the eggs produced by G5 and obtain the G6 generation. Screen larvae at the G6 stage for fluorescent markers.
   2. Discard larvae without fluorescent markers (Figure 4xi).
   3. Intercross mosquitoes that exhibit fluorescent markers.
   4. Continue this procedure for each subsequent generation until no non-fluorescent larva is observed. At this point, the mosquito line will be homozygous.

Results

Design and validation of gRNA-mediated gene targeting for HDR homology recombination
To ensure the desired gene is accurately targeted, we recommend selecting a couple of gRNAs and positioning the 5' and 3' homology arms close to the cutting site for HDR-mediated homologous recombination (Figure 1A). For example, we designed two gRNAs to target both sides of the start codon of the gene of interest and used the QF2-Hsp70-OpIE2-EC...

Discussion

CRISPR-Cas technology has changed the landscape of genome editing by promoting target-specific changes in chromosomes¹. Even though transposable elements were essential for the generation of the first transgenic mosquitoes, their insertion sites are somewhat random, and the expression of the cargo construct (promoter + gene) may not correspond to the expression profile of the actual gene due to a genome positional effect (i.e., insertion site), which usually leads to ectopic expression

Materials

Name	Company	Catalog Number	Comments
10x Cas9 reaction buffer	PNA Bio	CB01
Benchling software	Benchling	N/A	www.benchling.com
Cas9 dilution buffer	PNA Bio	CB03
Cas9 protein	PNA Bio	CP01-50
DH5α E. coli Competent Cells	New England Biolabs	C2987
Double-sided sticky tape	Scotch Permanent	3136
Drosophila vials	Genesee Scientific	32-109
Filter papers	GE Healthcare Life Science	1450-042
Fish food	Tetra	B00025Z6YI	goldfish flakes
Flugs	Genesee Scientific	AS273
Fluorescent microscope	Leica Microsystems	M165 FC
Gene fragment	Integrated DNA Technologies	N/A
gRNA	Synthego	N/A
Halocarbon oil 700	Sigma-Aldrich	H8898
Injection microscope	Leica Microsystems	DM2000
JM109  E. coli Competent Cells	Zymo Research	T3005
Microinjector	Eppendorf		FemtoJet 4x
Microloader Tips for Filling Femtotips	Eppendorf	E5242956003
Micromanipulator	Eppendorf		TransferMan 4r
Micropipette Pullers	Sutter Instrument	P-2000
Microscope Cover Glass	Fisherbrand	12-542-B
Microscope slide	Eisco	12-550-A3
Mouse blood (live mice used for feeding)	University of California	IACUC, Animal Use Protocol #S17187	Used for mosquito blood feeding; details comply with animal ethics protocols
NEB Q5 High-Fidelity DNA polymerase	New England Biolabs	M0491S
PCR Purification Kit	Qiagen	28004
Plasmid Miniprep Kit	Zymo Research	D4036
Quartz filament	Sutter Instruments	QF100-70-10
Transcription Clean-Up Kit	Fisher Scientific	AM1908
Ultra-pure water	Life Technologies	10977-023