Wir danken Dr. David O'Brochta und allen Mitgliedern des Insect Genetic Technologies Coordination Research Network für die Hilfe und Schulung, die sie uns und anderen bei der Implementierung genetischer Technologien bieten. Wir danken Channa Aluvihare besonders für die Optimierung des Mikromanipulationsprotokolls, damit Culex-Embryonen injiziert und geschlüpft werden können. Wir danken auch Devante Simmons und Joseph Urso, Studenten, die im Meuti-Labor arbeiten, für ihre Unterstützung bei der Pflege und dem Screening transgener Moskitos und Zora Elmkami von der ITF für die Unterstützung bei der Aufzucht und Vorbereitung von Moskitos für die Injektion. Diese Arbeit wurde durch ein Interdisciplinary Seeds Grant des Infectious Diseases Institute der OSU unterstützt, das MEM zur Verfügung gestellt wurde.

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<li>Mattingly, P. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+Culex+pipiens+complex%5BTitle%5D)+AND+%22Transactions+of+the+Royal+Entomological+Society+of+London%22%5BJournal%5D)">The Culex pipiens complex.</a> Transactions of the Royal Entomological Society of London. 102 (7), 331-342 (1951).</li>
<li>Eldridge, B. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+effect+of+temperature+and+photoperiod+on+blood-feeding+and+ovarian+development+in+mosquitoes+of+the+Culex+pipiens+complex%5BTitle%5D)+AND+%22The+American+Journal+of+Tropical+Medicine+and+Hygiene%22%5BJournal%5D)">The effect of temperature and photoperiod on blood-feeding and ovarian development in mosquitoes of the Culex pipiens complex.</a> The American Journal of Tropical Medicine and Hygiene. 17 (1), 133-140 (1968).</li>
<li>Spielman, A., Wong, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Environmental+control+of+ovarian+diapause+in+Culex+pipiens%5BTitle%5D)+AND+%22Annals+of+the+Entomological+Society+of+America%22%5BJournal%5D)">Environmental control of ovarian diapause in Culex pipiens.</a> Annals of the Entomological Society of America. 66 (4), 905-907 (1973).</li>
<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>
<li>Lin, S., Staahl, B. T., Alla, R. K., Doudna, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Enhanced+homology-directed+human+genome+engineering+by+controlled+timing+of+CRISPR%2FCas9+delivery%5BTitle%5D)+AND+%22eLife%22%5BJournal%5D)">Enhanced homology-directed human genome engineering by controlled timing of CRISPR/Cas9 delivery.</a> eLife. 3, 04766(2014).</li>
</ol>

RH arbeitet für die Insect Transformation Facility, die Dienstleistungen in der genetischen Veränderung von Insekten anbietet.

Dieses Protokoll stellt Methoden vor, um spezifische Mutationen in das Genom von Culex-Moskitos einzuführen und kann verwendet werden, um auch das Genom anderer Moskitos zu bearbeiten. Das Protokoll ist insoweit von Bedeutung, als es nicht nur spezifische Details zur Vorbereitung der Injektionsmaterialien enthält, sondern auch einen detaillierten Videoüberblick darüber, wie Moskitos dazu veranlasst werden, Eier zu legen, sowie wie diese Eier zubereitet und injiziert werden. Wir fassen auch zusammen, wie man...

Culex-Moskitos sind in den gemäßigten und tropischen Regionen der Welt verbreitet und übertragen mehrere tödliche Viren, darunter West-Nil-Virus1,St. Louis-Enzephalitis2 sowie filariale Nematoden, die Hundeherzwurm3 und Elephantiasis4verursachen. Mitglieder des Culex pipiens-Komplexes, zu dem Cx. quinquefasciatus, Cx. pipiens pipiens und Cx. pipiens molestusgehören , zeigen auffällige Variationen in vielen Aspekten ihrer Biologie. Zum Beispiel, während Cx. quinquefasciatus und Cx. pipiens molestus nicht in der Lage sind, in eine überwinternde Ruhephase5,6, Cx. pipiens pipiens zeigen robuste saisonale Reaktionen und treten in die Diapause als Reaktion auf kurze Tage7,8ein. Darüber hinaus neigen Cx. pipiens molestus dazu, anthropophiler zu sein, während Cx. pipiens und Cx. quinquefasciatus zoophiler sind6. In den Vereinigten Staaten und an vielen anderen Orten der Welt kreuzen sich diese Arten jedoch, was starke Auswirkungen auf die Krankheitsübertragung hat, da Hybriden der Cx. pipiens pipiens und Cx. pipiens molestus opportunistische Feeder sind und sowohl Vögel als auch Menschenbeißen 9und somit als Brückenvektoren für das West-Nil-Virus dienen. Das Studium dieser und anderer faszinierender Aspekte der Biologie von Culex-Moskitos wurde zum Teil behindert, weil Culex-Moskitos im Labor etwas schwieriger zu züchten sind als Aedes-Moskitos, die ruhe- und austrocknungsresistente Eier produzieren10 und weil funktionelle molekulare Werkzeuge für Culex-Arten nicht so gut entwickelt sind.
CRISPR / Cas9 Genom-Editing ist eine leistungsstarke Technologie, die verwendet wurde, um die Biologie mehrerer wichtiger Mückenarten11,12,13, einschließlich der Südlichen Hausmücke, Culex quinquefasciatus14,15,16. Diese von Jennifer Doudna und Emmanuelle Charpentier entwickelte Technologie nutzt eine natürliche bakterielle Abwehr gegen Viren durch bakteriell abgeleitete, CRISPR-assoziierte Endonukleasen (Cas-Proteine; siehe Review von Van der Oost et al.17). Bei der Injektion in tierische Embryonen können die Cas9-Proteine in Kombination mit einer geeigneten Leit-RNA doppelsträngige Brüche im Genom erzeugen. Dies geschieht am häufigsten durch die Verwendung des Cas9-Proteins, das mit Leit-RNAs komplexiert ist, die die Endonukleaseaktivität auf eine bestimmte Region des Genoms lenken. Nachdem das Cas9-Protein einen ortsspezifischen Doppelstrangbruch erzeugt hat, versucht die zelluläre Maschinerie, den Bruch mit einem von zwei Mechanismen zu reparieren. Die erste beinhaltet die Ligatur der beiden Enden durch nicht-homologe End-Joining (NHEJ), die fehleranfällig ist und oft Out-Frame-Insertionen und Deletionen im Genom erzeugt, die zu nicht-funktionellen Proteinen führen können, wodurch eine Knock-out-Mutation erzeugt wird. Alternativ kann die zelluläre Maschinerie die homologiegerichtete Reparatur (HDR) verwenden, indem sie ähnliche Sequenzen findet, um den Bruch korrekt zu reparieren. Die ähnliche Sequenz kann durch das zweite Chromosom im Organismus bereitgestellt werden (siehe Review18). Wenn die reparierte Sequenz jedoch genau mit der ursprünglichen Sequenz übereinstimmt, kann das Cas9-Protein die DNA erneut schneiden. Alternativ können Forscher auch ein Spenderplasmid einschließen, das homologe Sequenzen auf beiden Seiten der Schnittstelle der Zielsequenz mit einer alternativen Reparatursequenz enthält - oft ein fluoreszierendes Markerprotein, eine modifizierte Version des ursprünglichen Gens oder eine andere Modifikation -, die kopiert und in das Genom eingefügt oder "eingeklopft" werden kann.
Das Timing ist bei der Injektion von Embryonen entscheidend, und dies ist insbesondere der Fall, wenn CRISPR / Cas9-Genom-Editing verwendet wird, um Mutationen bei Insekten zu erzeugen. Dies liegt daran, dass das Cas9-Protein und gRNAs die größte Fähigkeit haben, Mutationen nur dann zu erzeugen, wenn sich der Embryo in seinem synzytialen Zustand befindet, bevor sich Zellmembranen gebildet haben und wenn mehrere Kerne im Embryo zugänglich sind. Bei Moskitos erreichen die Kerne die Peripherie ~ 2-4 Stunden nach der Eiablage, abhängig von der Temperatur19, und daher muss vor dieser Zeit eine erfolgreiche Mikroinjektion erfolgen. Darüber hinaus schneidet das Cas9-Protein jede Kern-DNA, auf die es zugreifen kann, so dass das Individuum, das aus der Injektion resultiert, ein Mosaik von Zellen enthält, von denen einige die gewünschte Mutation aufweisen und andere nicht. Damit diese Mutationen erfolgreich vererbt werden können, muss das Cas9-Protein DNA schneiden, die sich in der Keimbahn befindet und die zukünftigen Eier und Spermien hervorbringt. Um sicherzustellen, dass Mutationen in der Keimbahn erzeugt werden, ist es am besten, alle Materialien in der Nähe der Position der Polzellen im Embryo zu injizieren, die die Vorläufer der Insektenkeimbahn sind. Die Polzellen befinden sich in der Nähe des hinteren Endes der Culex-Embryonen 20. Neben der Injektion von Embryonen ist es unerlässlich, einen sorgfältigen Plan für die Kreuzung und das Screening von Nachkommen zu entwickeln, um die gewünschte Mutation zu erkennen.
Dieses Protokoll beschreibt, wie gRNAs erzeugt und mit Cas9-Protein komplexiert werden, um Injektionsmischungen herzustellen, sowie wie weibliche Moskitos von Culex pipiens dazu veranlasst werden, Eier zu legen und wie diese Eier für die CRISPR / Cas9-vermittelte Genombearbeitung vorbereitet und injiziert werden. Darüber hinaus beschreiben wir, wie injizierte Embryonen und ihre Nachkommen aufgezogen, gekreuzt und gescreent werden, um zu bestätigen, dass die gewünschte Mutation erhalten wurde. Mit diesem Protokoll generierten wir Nullmutationen für ein Gen von Interesse, Zyklus,im Buckeye-Stamm von Culex pipiens. Dieser Stamm wurde ursprünglich 2013 aus feldgesammelten Moskitos in Columbus, Ohio, entwickelt und wird vom Meuti-Labor gepflegt. Dieses Protokoll kann für zusätzliche Studien verwendet werden, die eine CRISPR/Cas9-Genombearbeitung bei Culex-Moskitos sowie anderen Mückenarten erfordern, und ist im Allgemeinen für die Verwendung von CRISPR/Cas9-Genom-Editing bei jeder Insektenart relevant.

<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-Moskitos sind die Hauptvektoren mehrerer Krankheiten, die sich negativ auf die Gesundheit von Mensch und Tier auswirken, einschließlich west-Nil-Virus und Krankheiten, die durch filariale Nematoden wie Hundeherzwurm und Elefantenasis verursacht werden. Vor kurzem wurde CRISPR / Cas9-Genom-Editing verwendet, um ortsgerichtete Mutationen zu induzieren, indem ein Cas9-Protein, das mit einer Leit-RNA (gRNA) komplexiert wurde, in frisch gelegte Embryonen mehrerer Insektenarten injiziert wurde, einschließlich Moskitos, die zu den Gattungen Anopheles und Aedesgehören. Die Manipulation und Injektion von Culex-Moskitos ist etwas schwieriger, da diese Moskitos ihre Eier aufrecht in Flößen ablegen und nicht einzeln wie andere Mückenarten. Hier beschreiben wir, wie man gRNAs entwirft, sie mit Cas9-Protein komplexiert, weibliche Moskitos von Culex pipiens dazu bringt, Eier zu legen, und wie man neu gelegte Embryonen für die Mikroinjektion mit Cas9 / gRNA vorbereitet und injiziert. Wir beschreiben auch, wie man injizierte Moskitos auf die gewünschte Mutation rückt und abschirmt. Die repräsentativen Ergebnisse zeigen, dass diese Technik verwendet werden kann, um ortsgerichtete Mutationen im Genom von Culex-Moskitos zu induzieren und mit leichten Modifikationen auch bei anderen Mückenarten Nullmutanten zu erzeugen.

Mit dem beschriebenen Protokoll konnten wir erfolgreich Embryonen von Cx. pipiensinjizieren und beobachteten eine hohe Überlebensrate unter den injizierten Embryonen (~ 55%, Abbildung 1). Frühere Studien hatten einen geringeren Prozentsatz des Überlebens, wahrscheinlich, weil der vordere Teil des Eifollikels am medizinischen Verbandsstreifen befestigt war, was verhinderte, dass Mückenlarven aus dem Chorion entweichen und erfolgreich ins Wasser schwammen. Die Sicherstellung, dass...

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 wird zunehmend zur Charakterisierung der Genfunktion in Nicht-Modellorganismen eingesetzt. Dieses Protokoll beschreibt, wie Man Knock-out-Linien von Culex pipienserzeugt, von der Vorbereitung von Injektionsmischungen über die Gewinnung und Injektion von Moskitoembryonen bis hin zur Aufzucht, Kreuzung und dem Screening injizierter Moskitos und ihrer Nachkommen auf gewünschte Mutationen.

Vorbereitung und Injektion von Embryonen von Culex-Moskitos zur Erzeugung von Nullmutationen mit 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

Research

JoVE Journal

Biology

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

6.9K Aufrufe.  The Ohio State University. CRISPR/Cas9 wird zunehmend zur Charakterisierung der Genfunktion in Nicht-Modellorganismen eingesetzt. Dieses Protokoll beschreibt, wie Man Knock-out-Linien von Culex pipienserzeugt, von der Vorbereitung von Injektionsmischungen über die Gewinnung und Injektion von Moskitoembryonen bis hin zur Aufzucht, Kreuzung und dem Screening injizierter Moskitos und ihrer Nachkommen auf gewünschte Mutationen.

Serie: 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.

Injektion eines. stephensi Embryonen zu generieren Malaria-resistenter Moskitos

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

Forschung

Biologie

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.

Mikroinjektion von A. aegypti Embryonen, um transgene Moskitos erhalten

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.

Keim Urzelle Transplantation für CRISPR/Cas9-basierte in Xenopus überspringen

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 ...

Genetik

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.

Embryo-Mikroinjektionstechniken für eine effiziente ortsspezifische Mutagenese bei 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, ...

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.

Sandfliege (Phlebotomus papatasi) Embryo Mikroinjektion für CRISPR/Cas9 Mutagenese

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.

Indel-Nachweis nach CRISPR/Cas9-Mutagenese mittels hochauflösender Schmelzeanalyse in der Mücke Aedes aegypti

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.

Ernte und Vorbereitung Drosophila Embryonen für die elektrophysiologische Ableitung und andere Verfahren

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.

Genome Editing bei der Gelbfiebermücke Aedes aegypti mittels CRISPR-Cas9

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.

Embryo-Mikroinjektion und Knockout-Mutanten-Identifizierung von CRISPR/Cas9 Genome-Edited Helicoverpa Armigera (Hübner)

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

Verhalten

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.

Mikroinjektion von Mückenembryonen: Eine Technik zur Verabreichung exogener DNA in das Eigelb des Embryos von Anopheles gambiae, um Keimbahnmutationen zu erzeugen