We thank Dr. David O&#8217;Brochta and all members of the Insect Genetic Technologies Coordination Research Network for the help and training that they provide to us and others on the implementation of genetic technologies. We especially thank Channa Aluvihare for optimizing the micromanipulation protocol to allow Culex embryos to be injected and hatch. We also thank Devante Simmons and Joseph Urso, undergraduate students working in the Meuti lab, for their assistance caring for and screening transgenic mosquitoes, and Zora Elmkami from the ITF for assistance rearing and prepping mosquitoes for injection. This work was supported by an Interdisciplinary Seeds grant from the Infectious Diseases Institute at OSU provided to MEM.

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RH works for the Insect Transformation Facility, which provides services in insect genetic modification.

This protocol presents methods to introduce specific mutations into the genome of Culex mosquitoes and can be used to edit the genome of other mosquitoes as well. The protocol is significant in that it provides specific details of not only how to prepare the injection materials, but also a detailed video overview of how to induce mosquitoes to lay eggs, as well as how to prepare and inject those eggs. We also summarize of how to take advantage of the biology of female Cx. pipiens to lay eggs in individual rafts and thereby screen a smaller proportion of offspring from each female for desired mutations. The methods presented here have been optimized for Cx. pipiens but can be adapted, with small adjustments, to edit the genomes of other mosquitoes or insects. Additionally, the micromanipulation and microinjection protocols described here are amenable to injecting several different materials into insect embryos, including transposons, dsRNA or other endonucleases such as TALENs or Zinc-Finger nucleases. Moreover, the beveling protocol generates microinjection needles with variable opening sizes therefore creating needles that can be used for injecting a wide variety of injection materials. The open size of the needle can be inferred by slowly decreasing the pressure of air after the needle has been beveled, and noting the pressure at which bubbles of air stop flowing from the tip of the newly opened needle. The less air pressure that is needed to produce bubbles indicates that the needle has a larger the opening size, and conversely, higher air pressures indicate that the needle has a smaller opening size. Needles with a larger opening sizes are better for injecting larger particles and more viscous materials, but will likely cause greater damage to the embryo and therefore reduce survival.
Microinjection experiments are in many ways a race against the clock. First, one must inject individual embryos within the first 2 hours of oviposition, before the chorion hardens and blastoderm formation occurs. Therefore, it is imperative to note when the mosquitoes were first placed in the oviposition chamber, how long it takes to manipulate the eggs for injection (we recommend no more than 20-30 minutes), and how long it takes to inject all embryos. Time is also limited when screening adult mosquitoes for mutations as Cx. pipiens are quite sensitive to their surroundings and mosquitoes generally only survive for 3-5 days in individual glass vials. Therefore, it is imperative to screen mosquito pupal exuvia as quickly as possible. Alternatively, one could also extract genomic DNA from a single mosquito leg14. In order to do so, however, the mosquitoes should first be anesthetized on ice. As we were weary of how the cold treatment and loss of limbs might impact mosquito survival and their ability to mate and lay viable eggs, we decided to instead screen the discarded pupal exuvia for mutations. The level of gDNA within the exuvia is likely lower than inside a mosquito leg, and this does add extra effort and time to separate mosquitoes at the pupal stage, but also ensures that all of the adults are unmated when they are released into their appropriate cages, which is absolutely vital. We feel that the results are worth it, but encourage others to consider if removing a leg might be a better course of action for their experimental needs.
The best way to expedite screening for mutations is to inject a plasmid containing a genetic marker, such as a fluorescent protein, flanked by homologous sequences on either side of the cut site. Rates of knock-in mutations using&#160;homology directed-repair (HDR) are generally lower than non-homologous end-joining (NHEJ), knock-out mutations (HDR=0.71%; NHEJ=24.87%;22). Therefore, the insertion of the marker will likely occur with a much lower frequency, but the time-savings offered by being able to quickly screen mosquito larvae under a fluorescent microscope may be worth the risk, depending on the project. Additionally, rates of HDR and knock-in mutations are enhanced when one also injects a plasmid containing the genetic sequence of the Cas9 protein, driven by the a well-characterized embryonic promotor, and the gRNA driven by the U6 promotor24,26. This is likely because early in embryonic development, when nuclei are rapidly dividing (S phase of the cell cycle), the error-prone but expedient NHEJ mechanism seems to be the preferred method of repairing double-stranded breaks (reviewed18). However, as embryogenesis progresses the cellular machinery is primed to use the more accurate HDR mechanism to repair double-stranded breaks (late S phase and G2 phase). Injecting a plasmid that contains the Cas9 sequence early in embryonic development allows the Cas9 protein to reach most target cells while the embryo is still in its syncytial state and before cellular membranes have formed, but the slight delay required to transcribe and translate the Cas9 protein seems to increase the likelihood that the endonucleasewill be active at a time when the developing embryo is more likely to use HDR to accurately repair double-stranded breaks in genomic DNA. For example, Lin et al.27 discovered that rates of HDR were enhanced to ~33% when the Cas9 protein complexed with gRNA were injected in human cell lines arrested in the M phase of the cell cycle. An additional benefit of delivering Cas9 and gRNAs on plasmids is that they are less viscous and therefore less likely to clog the needle during injections (R. Harrell, personal observation). However, until embyronic promoters are characterized and validated in Culex mosquitoes, we recommend injecting Cas9 protein or mRNA as described here.
Additionally, given the amount of time that one must dedicate to rearing and screening mosquitoes, we recommend having at least one full-time technician, student or researcher devoted to this task. We also recommend strategically considering how many null mutants one needs for a given experiment and the importance of genetic diversity within the null line. While we were able to identify mosquitoes in the F1 and F2 generations that had the mutation, only a handful of the F2 mosquitoes survived to adulthood the heterozygous mutant mosquitoes failed to produce viable offspring. We think that this is much less to do with the mutation, and more likely is a result of the lengthy period of time that both male and female mosquitoes were confined to glass tubes while we were screening them. This prolonged period of isolation most likely interfered with their ability to successfully mate and, in the case of females, obtain a blood meal and lay viable eggs. Outcrossing for a single generation and/or having optimized screening techniques in place should prevent this issue from occurring in the future. Therefore, by following this protocol we are confident that researchers will be able to successfully generate null lines of Culex mosquitoes and, with minor adjustments, additional insects.

Culex mosquitoes are distributed throughout the temperate and tropical regions of the world and transmit several deadly viruses including West Nile virus1, St. Louis encephalitis2 as well as filarial nematodes that cause canine heartworm3 and elephantiasis4. Members of the Culex pipiens complex, which includes Cx. quinquefasciatus, Cx. pipiens pipiens and Cx. pipiens molestus, show striking variations in many aspects of their biology. For example, while Cx. quinquefasciatus and Cx. pipiens molestus are incapable of entering an overwintering dormancy5,6, Cx. pipiens pipiens display robust seasonal responses and enter diapause in response to short days7,8. Additionally, Cx. pipiens molestus tend to be more anthropophilic while Cx. pipiens and Cx. quinquefasciatus are more zoophilic6. However, in the United States and throughout many other places in the world, these species interbreed, which has strong implications for disease transmission as hybrids of the Cx. pipiens pipiens and Cx. pipiens molestus are opportunistic feeders and will bite both birds and humans9, thereby serving as bridge vectors for West Nile virus. Studying these and other fascinating aspects of the biology of Culex mosquitoes has been hampered, in part, because Culex mosquitoes are slightly more difficult to rear in the lab than Aedes mosquitoes, which produce quiescent and desiccation-resistant eggs10 and because functional molecular tools are not as well developed for Culex species.
CRISPR/Cas9 genome editing is a powerful technology that has been used to evaluate the biology of several important mosquito species11,12,13, including the Southern house mosquito, Culex quinquefasciatus14,15,16. This technology, developed by Jennifer Doudna and Emmanuelle Charpentier, exploits a natural bacterial defense against viruses by bacterially-derived, CRISPR-associated endonucleases (Cas proteins; see review by Van der Oost et al.17). When injected into animal embryos, the Cas9 proteins in combination with an appropriate guide RNA can produce double-stranded breaks within the genome. This is most frequently done by using the Cas9 protein that is complexed with guide RNAs, which directs endonuclease activity to a specific region of the genome. After the Cas9 protein has created a site-specific double-stranded break, the cellular machinery attempts to repair the break using one of two mechanisms. The first entails ligating the two ends together through non-homologous end joining (NHEJ), which is error-prone and often produces out-frame insertions and deletions in the genome that can result in non-functional proteins, thereby generating a knock-out mutation. Alternatively, the cellular machinery might use homology-directed repair (HDR) by finding similar sequences to correctly repair the break. The similar sequence may be provided by the second chromosome within the organism (see review18). However, if the repaired sequence exactly matches the original sequence, the Cas9 protein will be able to again cut the DNA. Alternatively, researchers can also include a donor plasmid that contains homologous sequences on either side of the cut site of the target sequence with an alternative repair sequence&#8212;often a fluorescent marker protein, modified version of the original gene, or other modification&#8212;that can be copied and inserted into the genome, or &#8220;knocked-in.&#8221;
Timing is critical when injecting embryos, and this is especially the case when using CRISPR/Cas9 genome editing to create mutations in insects. This is because the Cas9 protein and gRNAs have the greatest capacity to generate mutations only when the embryo is in its syncytial state, before cellular membranes have formed and when multiple nuclei are accessible within the embryo. For mosquitoes, nuclei reach the periphery ~2-4 hours after oviposition, depending on temperature19, and therefore successful microinjection must occur before this time. Additionally, the Cas9 protein will cut any nuclear DNA that it can access, such that the individual resulting from the injection will contain a mosaic of cells, some having the desired mutation, and others not. In order for these mutations to be successfully inherited, the Cas9 protein must cut DNA that resides in the germline that will give rise to the future eggs and sperm. To ensure that mutations are generated in the germline it is best to inject all materials close to the location of the pole cells within the embryo, which are the progenitors of the insect germline. The pole cells are located near the posterior end of Culex embryos20. In addition to injecting embryos, it is imperative to develop a careful plan for crossing and screening offspring in order to detect the desired mutation.
This protocol describes how to generate gRNAs and complex them with Cas9 protein to prepare injection mixes, as well as how to induce female mosquitoes of Culex pipiens to lay eggs and how to prepare and inject those eggs for CRISPR/Cas9-mediated genome editing. Additionally, we describe how to rear, cross and screen injected embryos and their progeny to confirm that the desired mutation has been obtained. Using this protocol, we generated null mutations for a gene of interest, cycle, in the Buckeye strain of Culex pipiens. This strain was originally established in 2013 from field-collected mosquitoes in Columbus, Ohio and is maintained by the Meuti lab. This protocol can be used for additional studies that require CRISPR/Cas9 genome editing in Culex mosquitoes, as well as other mosquito species, and, more generally, is relevant to employing CRISPR/Cas9 genome editing to any insect species.

<table><tbody><tr><td>Artificial Membrane Feeder</td><td>Hemotek</td><td>SP5W1-3</td><td>Company location: Blackburn, UK</td></tr><tr><td>ATP</td><td>Invitrogen</td><td>18330019</td><td>Company location: Carlsbad, CA, USA</td></tr><tr><td>Borosilicate glass mirocapillary tubes, 1 mm outer diameter</td><td>World Precision Instruments</td><td>1B100-6</td><td>Company Location: Sarasota, FL, USA</td></tr><tr><td>BV10 Needle Beveler</td><td>Sutter Instruments</td><td>BV-10-B</td><td>Company Location: Nobato, CA, USA</td></tr><tr><td>Whatman Circular filter paper (12.5 cm)</td><td>Sigma Aldrich</td><td>WHA1001125</td><td>Company Location: St. Louis, MO, USA</td></tr><tr><td>Conical tube (50 mL)</td><td>Thermo Fisher Scientific</td><td>339652</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>Fisherbrand course filter paper with fast flow rate</td><td>Thermo Fisher Scientific</td><td>09-800</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>Cover glass (24 x 40 mm)</td><td>Thermo Fisher Scientific</td><td>50-311-20</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>Dental dam</td><td>Henry Schein Inc</td><td>1010171</td><td>Company Location: Melville, NY USA</td></tr><tr><td>Scotch double-sided tape</td><td>Thermo Fisher Scientific</td><td>NC0879005</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>FemtoJet 4i microinjector</td><td>Eppendorf</td><td>5252000021</td><td>Company Location: Hamburg, Germany</td></tr><tr><td>Glass vial (2 dram)</td><td>Thermo Fisher Scientific</td><td>033401C</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>Halocarbon oil</td><td>Sigma Aldrich</td><td>H8898-50ML</td><td>Company Location: St. Louis, MO, USA</td></tr><tr><td>P-2000 Laser Needle Puller</td><td>Sutter Instruments</td><td>P-2000/G</td><td>Company Location: Nobato, CA, USA</td></tr><tr><td>Parafilm</td><td>Thermo Fisher Scientific</td><td>50-998-944</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>PC-100 Weighted Needle Puller</td><td>Narishige</td><td>PC-100</td><td>This is compatabile with the earlier PC-10 model, which has been discontinued. Company Location: Amityville, NY, USA</td></tr><tr><td>Phire Direct PCR Kit</td><td>Thermo Fisher Scientific</td><td>F140WH</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>Kodak Photo-Flo (1%)</td><td>Thermo Fisher Scientific</td><td>50-268-05</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>Quartz glass mirocapillary tubes, 1 mm outer diameter</td><td>Capillary Tube Supplies Limited</td><td>QGCT 1.0</td><td>Company Location: Cornwall, UK</td></tr><tr><td>Guide-it&amp;trade; sgRNA Screening Kit</td><td>Takara, Bio USA</td><td>632639</td><td>This kit allows you to determine if gRNAs cut DNA sequences in vitro. Company Location: Mountain View, CA, USA</td></tr><tr><td>Sigmacote</td><td>Sigma Aldrich</td><td>SL2-100ML</td><td>Company Location: St. Louis, MO, USA</td></tr><tr><td>Small petri dishes (35X10 mm)</td><td>Thermo Fisher Scientific</td><td>50-190-0273</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>Sodium citrate chicken blood</td><td>Lampire biologicals</td><td>7201406</td><td>Company Location: Everett, PA, USA</td></tr><tr><td>Fisherbrand Square petri dish (10 cm x 10 cm)</td><td>Thermo Fisher Scientific</td><td>FB0875711A</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>Tegaderm</td><td>Henry Schein Inc.</td><td>7771180</td><td>Company Location: Melville, NY USA</td></tr><tr><td>Tropical fish food</td><td>Tetramin</td><td>N/A</td><td></td></tr><tr><td>Whatman filter paper</td><td>Thermo Fisher Scientific</td><td>09-927-826</td><td>Company Location: Waltham, MA, USA</td></tr><tr><td>Whatman filter paper, 4.25 cm</td><td>Sigma Aldrich</td><td>1001-042</td><td>Company Location: St. Louis, MO, USA</td></tr></tbody></table>

preparing and injecting embryos of culex mosquitoes to generate null mutations using crispr/cas9

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by filarial nematodes such as canine heartworm and elephantasis. Recently, CRISPR/Cas9 genome editing has been used to induce site-directed mutations by injecting a Cas9 protein that has been complexed with a guide RNA (gRNA) into freshly laid embryos of several insect species, including mosquitoes that belong to the genera Anopheles and Aedes. Manipulating and injecting Culex mosquitoes is slightly more difficult as these mosquitoes lay their eggs upright in rafts rather than individually like other species of mosquitoes. Here we describe how to design gRNAs, complex them with Cas9 protein, induce female mosquitoes of Culex pipiens to lay eggs, and how to prepare and inject newly laid embryos for microinjection with Cas9/gRNA. We also describe how to rear and screen injected mosquitoes for the desired mutation. The representative results demonstrate that this technique can be used to induce site-directed mutations in the genome of Culex mosquitoes and, with slight modifications, can be used to generate null-mutants in other mosquito species as well.&#160;

Using the described protocol, we were able to successfully inject embryos of Cx. pipiens, and observed a high rate of survival among the injected embryos (~55%, Figure 1). Earlier trials had a lower percentage of survival, likely because the anterior of the egg follicle was attached to the medical dressing strip, preventing mosquito larvae from escaping from the chorion and successfully swimming into the water. Ensuring that the anterior end extends beyond the strip of medical dressing greatly increased larval survival, and results in high quality offspring that are capable of developing to adulthood and reproducing (Figure 2B).
Subsequent experiments and screening indicate that the null mutation for the circadian clock gene cycle (cyc; KM355981) made it into the germline of the injected embryos (Figure 3). Given that our primers amplified a large region of the cyc gene near the cut-sites, it was difficult to observe two separate bands in heterozygous, F1 mosquitoes. This demonstrates the utility of designing primers that will amplify fragments that are ~100-200 bp around the cut site, and also the importance of sequencing all suspected mutant mosquitoes. Sequencing revealed that approximately 10% of the screened mosquitoes showed a small insertion or deletion near the Cas9 cut site of the first guide RNA that we had designed (Figure 3), suggesting that this was a highly effective gRNA and that we successfully targeted the pole cells during microinjections. The F1 individuals successfully mated and produced viable offspring (F2 generation), some of which also contained the mutation.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61651/61651fig1.jpg" /> 
Figure 1: Example of a beveled borosilicate needle. This needle was fabricated from a siliconized borosilicate microcapillary tube, pulled using a PC-10 vertical needle puller, and then beveled on a BV-10 Beveller. The needle is viewed under ~63x magnification. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61651/61651fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61651/61651fig2.jpg" /> 
Figure 2: Survival of the F0&#173; generation across their development. (A) Image of 1st instar larvae that hatched from 3 slides containing injected embryos. (B) Graphical representation of the number of individuals at each life stage. Of the 801 embryos that were injected, 441 1st instar larvae hatched, and of these 149 individuals reached pupation, resulting in a total of 121 adults (73 males and 43 females). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61651/61651fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61651/61651fig3.jpg" /> 
Figure 3: Representative results showing the transmission of the desired mutation. The top two sequences represent the sequence of the cycle gene in Culex quinquefascaitus (Cx.quinq; XP_001865023.1) and in wild type females of Cx. pipiens (WT.F; KM355981). The sequences below represent sequences in three male F&#173;1 offspring (I.DM1; II.JM4; II.K10M). The green box represents the PAM sequence recognized by the Cas9 protein (CGG) while the green arrow represents where the Cas9 protein cleaved genomic DNA. These results show that short deletions of 2 or 4 nucleotides were inherited in the F1 males. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61651/61651fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9 at JoVE.com

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

CRISPR/Cas9 is increasingly used to characterize gene function in non-model organisms. This protocol describes how to generate knock-out lines of Culex pipiens, from preparing injection mixes, to obtaining and injecting mosquito embryos, as well as how to rear, cross, and screen injected mosquitoes and their progeny for desired mutations.

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by ...

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6.9K Views.  The Ohio State University. CRISPR/Cas9 is increasingly used to characterize gene function in non-model organisms. This protocol describes how to generate knock-out lines of Culex pipiens, from preparing injection mixes, to obtaining and injecting mosquito embryos, as well as how to rear, cross, and screen injected mosquitoes and their progeny for desired mutations.

Video: Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

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Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

Genes essential for survival pose technical hurdles for creating mutant lines. Leapfrogging circumvents lethality by combining genome editing with primordial germ cell transplantation to create wild-type animals carrying germline mutations. Leapfrogging also permits the efficient generation of homozygous null mutants in the F1 generation. Here, the transplantation step is demonstrated.

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Genetics

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, Eastern equine encephalitis, lymphatic filariasis, and Saint Louis encephalitis. Notably, avian malaria has played a major role in the extinction of numerous endemic island bird species, while WNV has become an important vector-borne disease in the United States. To gain further insight into C. quinquefasciatus biology and expand their genetic control toolbox, we need to develop more efficient and affordable methods for genome engineering in this species. However, some biological traits unique to Culex mosquitoes, particularly their egg rafts, have made it difficult to perform microinjection procedures required for genome engineering. To address these challenges, we have developed an optimized embryo microinjection protocol that focuses on mitigating the technical obstacles associated with the unique characteristics of Culex mosquitoes. These procedures demonstrate optimized methods for egg collection, egg raft separation and other handling procedures essential for successful microinjection in C. quinquefasciatus. When coupled with the CRISPR/Cas9 genome editing technology, these procedures allow us to achieve site-specific, efficient and heritable germline mutations, which are required to perform advanced genome engineering and develop genetic control technologies in this important, but currently understudied, disease vector.

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Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to visceral pathology. Deciphering the nature of the vector/parasite interactions is of primary importance for better understanding of Leishmania transmission to their hosts. Among the parameters controlling the sand fly vector competence (i.e. their ability to carry and transmit pathogens), parameters intrinsic to these insects were shown to play a key role. Insect immune response, for example, impacts sand fly vector competence to Leishmania. The study of such parameters has been limited by the lack of methods of gene expression modification adapted for use in these non-model organisms. Gene downregulation by small interfering RNA (siRNA) is possible, but in addition to being technically challenging, the silencing leads to only a partial loss of function, which cannot be transmitted from generation to generation. Targeted mutagenesis by CRISPR/Cas9 technology was recently adapted to the Phlebotomus papatasi sand fly. This technique leads to the generation of transmissible mutations in a specifically chosen locus, allowing to study the genes of interest. The CRISPR/Cas9 system relies on the induction of targeted double-strand DNA breaks, later repaired by either Non-Homologous End Joining (NHEJ) or by Homology Driven Repair (HDR). NHEJ consists of a simple closure of the break and frequently leads to small insertion/deletion events. In contrast, HDR uses the presence of a donor DNA molecule sharing homology with the target DNA as a template for repair. Here, we present a sand fly embryo microinjection method for targeted mutagenesis by CRISPR/Cas9 using NHEJ, which is the only genome modification technique adapted to sand fly vectors to date.

Sand Fly Phlebotomus papatasi Embryo Microinjection for CRISPR/Cas9 Mutagenesis

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Mosquito gene editing has become routine in several laboratories with the establishment of systems such as transcription-activator-like effector nucleases (TALENs), zinc-finger nucleases (ZFNs), and homing endonucleases (HEs). More recently, clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) technology has offered an easier and cheaper alternative for precision genome engineering. Following nuclease action, DNA repair pathways will fix the broken DNA ends, often introducing indels. These out-of-frame mutations are then used for understanding gene function in the target organisms. A drawback, however, is that mutant individuals carry no dominant marker, making identification and tracking of mutant alleles challenging, especially at scales needed for many experiments. 
High-resolution melt analysis (HRMA) is a simple method to identify variations in nucleic acid sequences and utilizes PCR melting curves to detect such variations. This post-PCR analysis method uses fluorescent double-stranded DNA-binding dyes with instrumentation that has temperature ramp control data capture capability and is easily scaled to 96-well plate formats. Described here is a simple workflow using HRMA for the rapid detection of CRISPR/Cas9-induced indels and the establishment of mutant lines in the mosquito Ae. aegypti. Critically, all steps can be performed with a small amount of leg tissue and do not require sacrificing the organism, allowing genetic crosses or phenotyping assays to be performed after genotyping.

Indel Detection following CRISPR/Cas9 Mutagenesis using High-resolution Melt Analysis in the Mosquito Aedes aegypti

This article details a protocol for rapid identification of indels induced by CRISPR/Cas9 and selection of mutant lines in the mosquito Aedes aegypti using high-resolution melt analysis.

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Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

This technique exposes the Drosophila embryonic neuromusculature for immunohistochemistry or electrophysiological recording. It is useful for studying early events in neuromuscular development or performing electrophysiology in mutants that cannot hatch.

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

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.

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

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Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera (H&#252;bner)

The cotton bollworm, Helicoverpa armigera, is one of the most destructive pests in the world. A combination of molecular genetics, physiology, functional genomics, and behavioral studies has made H. armigera a model species in Lepidoptera Noctuidae. To study the in vivo functions of and interactions between different genes, clustered regularly interspaced short palindromic repeats (CRISPR)/ associated protein 9 (Cas9) genome editing technology is a convenient and effective method used for performing functional genomic studies. In this study, we provide a step-by-step systematic method to complete gene knockout in H. armigera using the CRISPR/Cas9 system. The design and synthesis of guide RNA (gRNA) are described in detail. Then, the subsequent steps consisting of gene-specific primer design for guide RNA (gRNA) creation, embryo collection, microinjection, insect rearing, and mutant detection are summarized. Finally, troubleshooting advice and notes are provided to improve the efficiency of gene editing. Our method will serve as a reference for the application of CRISPR/Cas9 genome editing in H. armigera as well as other Lepidopteran moths.

Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera H&#252;bner

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The cotton bollworm, Helicoverpa armigera, is one of the most destructive pests in the world. A combination of molecular genetics, physiology, functional ...

Behavior

Mosquito Embryo Microinjection: A Technique to Deliver Exogenous DNA into Embryo Yolk of Anopheles Gambiae to Generate Germline Mutations

Use a micro-loader tip to fill the needles with two microliters of DNA mixture. Insert the needle into the needle-holder, and connect the automated pressure pump tubing. Align the needle so that it makes an angle of 15 degrees with the slide&#39;s plane.
Open the needle tip by carefully touching the first egg. Then, insert the needle&#8217;s tip to about 10 micrometers in the posterior pole. A successful injection will lead to a small movement of the cytoplasm within the egg.
Use the microscope co-axial stage controls to move to the next egg to continue injection. To ensure that the needle tip remains open and has not been clogged, press the &#39;Inject&#39; button before entering another embryo, and visualize the small droplet at the opening of the needle. If the needle gets clogged, press the &#39;Clear&#39; button to clear the needle, and repeat the droplet visualization test.
Adjust the pressure as needed, if the needle tip opening gets slightly bigger to ensure that the droplet size stays small. Inject about 40 to 50 eggs with one needle. After injections are complete, rinse the eggs off into a glass container lined with filter paper and filled with 50 milliliters of deionized water.

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