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.

<ol><li>Hamer, G. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Culex+pipiens+%28Diptera%3A+Culicidae%29%3A+a+bridge+vector+of+West+Nile+virus+to+humans%5BTitle%5D)+AND+%22Journal+of+Medical+Entomology%22%5BJournal%5D)">Culex pipiens (Diptera: Culicidae): a bridge vector of West Nile virus to humans.</a> Journal of Medical Entomology. 45 (1), 125-128 (2008).</li>
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<li>Fritz, M. L., Walker, E. D., Miller, J. R., Severson, D. W., Dworkin, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Divergent+host+preferences+of+above-and+below-ground+Culex+pipiens+mosquitoes+and+their+hybrid+offspring%5BTitle%5D)+AND+%22Medical+and+Veterinary+Entomology%22%5BJournal%5D)">Divergent host preferences of above-and below-ground Culex pipiens mosquitoes and their hybrid offspring.</a> Medical and Veterinary Entomology. 29 (2), 115-123 (2015).</li>
<li>Meola, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+influence+of+temperature+and+humidity+on+embryonic+longevity+in+Aedes+aegypti%5BTitle%5D)+AND+%22Annals+of+the+Entomological+Society+of+America%22%5BJournal%5D)">The influence of temperature and humidity on embryonic longevity in Aedes aegypti.</a> Annals of the Entomological Society of America. 57 (4), 468-472 (1964).</li>
<li>Dong, S., Lin, J., Held, N. L., Clem, R. J., Passarelli, A. L., Franz, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Heritable+CRISPR%2FCas9-mediated+genome+editing+in+the+yellow+fever+mosquito%2C+Aedes+aegypti%5BTitle%5D)+AND+%22PloS+one%22%5BJournal%5D)">Heritable CRISPR/Cas9-mediated genome editing in the yellow fever mosquito, Aedes aegypti.</a> PloS one. 10 (3), (2015).</li>
<li>Kyrou, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+CRISPR-Cas9+gene+drive+targeting+doublesex+causes+complete+population+suppression+in+caged+Anopheles+gambiae+mosquitoes%5BTitle%5D)+AND+%22Nature+Biotechnology%22%5BJournal%5D)">A CRISPR-Cas9 gene drive targeting doublesex causes complete population suppression in caged Anopheles gambiae mosquitoes.</a> Nature Biotechnology. 36 (11), 1062-1066 (2018).</li>
<li>Liu, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Construction+of+an+efficient+genomic+editing+system+with+CRISPR%2FCas9+in+the+vector+mosquito+Aedes+albopictus%5BTitle%5D)+AND+%22Insect+Science%22%5BJournal%5D)">Construction of an efficient genomic editing system with CRISPR/Cas9 in the vector mosquito Aedes albopictus.</a> Insect Science. 26 (6), 1045-1054 (2019).</li>
<li>Itokawa, K., Komagata, O., Kasai, S., Ogawa, K., Tomita, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Testing+the+causality+between+CYP9M10+and+pyrethroid+resistance+using+the+TALEN+and+CRISPR%2FCas9+technologies%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">Testing the causality between CYP9M10 and pyrethroid resistance using the TALEN and CRISPR/Cas9 technologies.</a> Scientific Reports. 6, 24652(2016).</li>
<li>Li, M., Li, T., Liu, N., Raban, R. R., Wang, X., Akbari, O. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Methods+for+the+generation+of+heritable+germline+mutations+in+the+disease+vector+Culex+quinquefasciatus+using+clustered+regularly+interspaced+short+palindrome+repeats-associated+protein+9%5BTitle%5D)+AND+%22Insect+Molecular+Biology%22%5BJournal%5D)">Methods for the generation of heritable germline mutations in the disease vector Culex quinquefasciatus using clustered regularly interspaced short palindrome repeats-associated protein 9.</a> Insect Molecular Biology. 29, 214-220 (2019).</li>
<li>Anderson, M. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((CRISPR%2FCas9+gene+editing+in+the+West+Nile+Virus+vector%2C+Culex+quinquefasciatus+Say%5BTitle%5D)+AND+%22PloS+one%22%5BJournal%5D)">CRISPR/Cas9 gene editing in the West Nile Virus vector, Culex quinquefasciatus Say.</a> PloS one. 14 (11), (2019).</li>
<li>Van der Oost, J., Jore, M. M., Westra, E. R., Lundgren, M., Brouns, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((CRISPR-based+adaptive+and+heritable+immunity+in+prokaryotes%5BTitle%5D)+AND+%22Trends+in+Biochemical+Sciences%22%5BJournal%5D)">CRISPR-based adaptive and heritable immunity in prokaryotes.</a> Trends in Biochemical Sciences. 34 (8), 401-407 (2009).</li>
<li>Heyer, W. D., Ehmsen, K. T., Liu, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Regulation+of+homologous+recombination+in+eukaryotes%5BTitle%5D)+AND+%22Annual+Review+of+Genetics%22%5BJournal%5D)">Regulation of homologous recombination in eukaryotes.</a> Annual Review of Genetics. 44, 113-139 (2010).</li>
<li>Monnerat, A. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Anopheles+albitarsis+embryogenesis%3A+morphological+identification+of+major+events%5BTitle%5D)+AND+%22Memorias+do+Instituto+Oswaldo+Cruz%22%5BJournal%5D)">Anopheles albitarsis embryogenesis: morphological identification of major events.</a> Memorias do Instituto Oswaldo Cruz. 97 (4), 589-596 (2002).</li>
<li>Davis, C. W. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+comparative+study+of+larval+embryogenesis+in+the+mosquito+Culex+fatigans+Wiedemann+%28Diptera%3A+Culicidae%29+and+the+sheep-fly+Lucilia+sericata+Meigen+%28Diptera%3A+Calliphoridae%29%5BTitle%5D)+AND+%22Australian+Journal+of+Zoology%22%5BJournal%5D)">A comparative study of larval embryogenesis in the mosquito Culex fatigans Wiedemann (Diptera: Culicidae) and the sheep-fly Lucilia sericata Meigen (Diptera: Calliphoridae).</a> Australian Journal of Zoology. 15 (3), 547-579 (1967).</li>
<li>Port, F., Chen, H. M., Lee, T., Bullock, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Optimized+CRISPR%2FCas+tools+for+efficient+germline+and+somatic+genome+engineering+in+Drosophila%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences%22%5BJournal%5D)">Optimized CRISPR/Cas tools for efficient germline and somatic genome engineering in Drosophila.</a> Proceedings of the National Academy of Sciences. 111 (29), 2967-2976 (2014).</li>
<li>Kistler, K. E., Vosshall, L. B., Matthews, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Genome+engineering+with+CRISPR-Cas9+in+the+mosquito+Aedes+aegypti%5BTitle%5D)+AND+%22Cell+Reports%22%5BJournal%5D)">Genome engineering with CRISPR-Cas9 in the mosquito Aedes aegypti.</a> Cell Reports. 11 (1), 51-60 (2015).</li>
<li>Gokcezade, J., Sienski, G., Duchek, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Efficient+CRISPR%2FCas9+plasmids+for+rapid+and+versatile+genome+editing+in+Drosophila%5BTitle%5D)+AND+%22G3%3A+Genes%2C+Genomes%2C+Genetics%22%5BJournal%5D)">Efficient CRISPR/Cas9 plasmids for rapid and versatile genome editing in Drosophila.</a> G3: Genes, Genomes, Genetics. 4 (11), 2279-2282 (2014).</li>
<li>Hwang, W. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Efficient+in+vivo+genome+editing+using+RNA-guided+nucleases%5BTitle%5D)+AND+%22Nature+Biotechnology%22%5BJournal%5D)">Efficient in vivo genome editing using RNA-guided nucleases.</a> Nature Biotechnology. 31 (3), 227-229 (2013).</li>
<li>Basu, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Silencing+of+end-joining+repair+for+efficient+site-specific+gene+insertion+after+TALEN%2FCRISPR+mutagenesis+in+Aedes+aegypti%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences%22%5BJournal%5D)">Silencing of end-joining repair for efficient site-specific gene insertion after TALEN/CRISPR mutagenesis in Aedes aegypti.</a> Proceedings of the National Academy of Sciences. 112 (13), 4038-4043 (2015).</li>
<li>Hammond, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+CRISPR-Cas9+gene+drive+system+targeting+female+reproduction+in+the+malaria+mosquito+vector+Anopheles+gambiae%5BTitle%5D)+AND+%22Nature+Biotechnology%22%5BJournal%5D)">A CRISPR-Cas9 gene drive system targeting female reproduction in the malaria mosquito vector Anopheles gambiae.</a> Nature Biotechnology. 34 (1), 78-83 (2016).</li>
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</ol>

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 individu...

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 dress...

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 ...

preparing-injecting-embryos-culex-mosquitoes-to-generate-null

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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

Injection of An. stephensi Embryos to Generate Malaria-resistant Mosquitoes

The introduction of exogenous genes into the genomes of mosquitoes requires microinjection techniques tailored to the specific species of interest. This video protocol demonstrates a method used by the James laboratory to microinject DNA constructs into Anopheles stephensi embryos for the generation of transformed mosquitoes. Techniques for preparing microinjection needles, collecting and preparing embryos and performing the microinjection are illustrated.

Anopheles stephensi mosquitoes are vectors for malaria inhabiting India and throughout Asia. This video demonstrates the technique for performing microinjections of this species with transgenes that will confer resistance to the malaria to the mosquito. Much of the methodology demonstrated in this video is applicable to microinjection techniques of other mosquito species.

The introduction of exogenous genes into the genomes of mosquitoes requires microinjection techniques tailored to the specific species of interest. This ...

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever.  The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever. The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue ...

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

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the African clawed frog, Xenopus, a longstanding model organism for biomedical research. Traditional breeding schemes for creating homozygous mutant lines with CRISPR/Cas9-targeted mutagenesis have several time-consuming and laborious steps. To facilitate the creation of mutant embryos, particularly to overcome the obstacles associated with knocking out genes that are essential for embryogenesis, a new method called leapfrogging was developed. This technique leverages the robustness of Xenopus embryos to &#34;cut and paste&#34; embryological methods. Leapfrogging utilizes the transfer of primordial germ cells (PGCs) from efficiently-mutagenized donor embryos into PGC-ablated wildtype siblings. This method allows for the efficient mutation of essential genes by creating chimeric animals with wildtype somatic cells that carry a mutant germline. When two F0 animals carrying &#34;leapfrog transplants&#34; (i.e., mutant germ cells) are intercrossed, they produce homozygous, or compound heterozygous, null F1 embryos, thus saving a full generation time to obtain phenotypic data. Leapfrogging also provides a new approach for analyzing maternal effect genes, which are refractory to F0 phenotypic analysis following CRISPR/Cas9 mutagenesis. This manuscript details the method of leapfrogging, with special emphasis on how to successfully perform PGC transplantation.

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.

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the ...

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.

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

This protocol describes microinjection procedures for Culex quinquefasciatus embryos that are optimized to work with CRISPR/Cas9 gene editing tools. This technique can efficiently generate site-specific, heritable, germline mutations that can be used for building genetic technologies in this understudied disease vector.

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, ...

Sand Fly (Phlebotomus papatasi) Embryo Microinjection for CRISPR/Cas9 Mutagenesis

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

This protocol details the steps of CRISPR/Cas9 targeted mutagenesis in sand flies: embryo collection, injection, insect rearing, and identification as well as selection of mutations of interest.

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to ...

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

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.

Mosquito gene editing has become routine in several laboratories with the establishment of systems such as transcription-activator-like effector nucleases ...

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.

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

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.

The emergence of the clustered, regularly interspersed, short palindromic repeats (CRISPR)-Cas9 technology has revolutionized the genetic engineering ...

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

Presented here is a protocol of Helicoverpa armigera (H&#252;bner) embryo microinjection and knockout mutant identification created by CRISPR/Cas9 genome editing. Mutant insects enable further research of gene function and interaction among different genes in vivo.

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.

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

Summary

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