Name: sand fly (phlebotomus papatasi) embryo microinjection for crispr/cas9 mutagenesis
Uploaded: 2020-11-17T15:00:00
Description: Watch this Scientific Journal Video about Sand Fly Phlebotomus papatasi Embryo Microinjection for CRISPR/Cas9 Mutagenesis at JoVE.com

The authors thank Vanessa Meldener-Harrell for critical reading of the manuscript.

The use of mice as a source of blood for sand fly feeding was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health (NIH). The protocol was approved by the Animal Care and Use Committee of the NIAID, NIH (protocol number LPD 68E). Invertebrates are not covered under NIH guidelines.
1. Needle preparation (Figure 1)
<ol>
	<li>Pull needles and bevel as described in Meut...

<ol>
	<li>Lawyer, P., Killick-Kendrick, M., Rowland, T., Rowton, E., Volf, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laboratory+colonization+and+mass+rearing+of+phlebotomine+sand+flies+(Diptera,+Psychodidae).">Laboratory colonization and mass rearing of phlebotomine sand flies (Diptera, Psychodidae).</a> Parasite. 24, 42 (2017).</li><li>Pelletier, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Specific+recognition+of+Leishmania+major+poly-beta-galactosyl+epitopes+by+galectin-9:+possible+implication+of+galectin-9+in+interaction+between+L.+major+and+host+cells.">Specific recognition of Leishmania major poly-beta-galactosyl epitopes by galectin-9: possible implication of galectin-9 in interaction between L. major and host cells.</a> Journal Biological Chemistry. 278 (25), 22223-22230 (2003).</li><li>Kamhawi, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+role+for+insect+galectins+in+parasite+survival.">A role for insect galectins in parasite survival.</a> Cell. 119 (3), 329-341 (2004).</li><li>Louradour, I., Ghosh, K., Inbar, E., Sacks, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9+Mutagenesis+in+Phlebotomus+papatasi:+the+Immune+Deficiency+Pathway+Impacts+Vector+Competence+for+Leishmania+major.">CRISPR/Cas9 Mutagenesis in Phlebotomus papatasi: the Immune Deficiency Pathway Impacts Vector Competence for Leishmania major.</a> MBio. 10 (4), (2019).</li><li>Xi, Z., Ramirez, J. L., Dimopoulos, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Aedes+aegypti+toll+pathway+controls+dengue+virus+infection.">The Aedes aegypti toll pathway controls dengue virus infection.</a> PLoS Pathogens. 4 (7), 1000098 (2008).</li><li>Ramirez, J. L., Dimopoulos, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Toll+immune+signaling+pathway+control+conserved+anti-dengue+defenses+across+diverse+Ae.+aegypti+strains+and+against+multiple+dengue+virus+serotypes.">The Toll immune signaling pathway control conserved anti-dengue defenses across diverse Ae. aegypti strains and against multiple dengue virus serotypes.</a> Development and Comparative Immunology. 34 (6), 625-629 (2010).</li><li>Ramirez, J. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reciprocal+tripartite+interactions+between+the+Aedes+aegypti+midgut+microbiota,+innate+immune+system+and+dengue+virus+influences+vector+competence.">Reciprocal tripartite interactions between the Aedes aegypti midgut microbiota, innate immune system and dengue virus influences vector competence.</a> PLoS Neglected Tropical Diseases. 6 (3), 1561 (2012).</li><li>Hu, C., Aksoy, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Innate+immune+responses+regulate+trypanosome+parasite+infection+of+the+tsetse+fly+Glossina+morsitans+morsitans.">Innate immune responses regulate trypanosome parasite infection of the tsetse fly Glossina morsitans morsitans.</a> Molecular Microbiology. 60 (5), 1194-1204 (2006).</li><li>Meister, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anopheles+gambiae+PGRPLC-mediated+defense+against+bacteria+modulates+infections+with+malaria+parasites.">Anopheles gambiae PGRPLC-mediated defense against bacteria modulates infections with malaria parasites.</a> PLoS Pathogens. 5 (8), 1000542 (2009).</li><li>Meister, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immune+signaling+pathways+regulating+bacterial+and+malaria+parasite+infection+of+the+mosquito+Anopheles+gambiae.">Immune signaling pathways regulating bacterial and malaria parasite infection of the mosquito Anopheles gambiae.</a> Proceedings of the National Academy of Sciences U S A. 102 (32), 11420-11425 (2005).</li><li>Telleria, E. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Caspar-like+gene+depletion+reduces+Leishmania+infection+in+sand+fly+host+Lutzomyia+longipalpis.">Caspar-like gene depletion reduces Leishmania infection in sand fly host Lutzomyia longipalpis.</a> Journal of Biological Chemistry. 287 (16), 12985-12993 (2012).</li><li>Sant'Anna, M. R., Alexander, B., Bates, P. A., Dillon, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+silencing+in+phlebotomine+sand+flies:+Xanthine+dehydrogenase+knock+down+by+dsRNA+microinjections.">Gene silencing in phlebotomine sand flies: Xanthine dehydrogenase knock down by dsRNA microinjections.</a> Insect Biochemistry and Molecular Biology. 38 (6), 652-660 (2008).</li><li>Sant'anna, M. R., Diaz-Albiter, H., Mubaraki, M., Dillon, R. J., Bates, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhibition+of+trypsin+expression+in+Lutzomyia+longipalpis+using+RNAi+enhances+the+survival+of+Leishmania.">Inhibition of trypsin expression in Lutzomyia longipalpis using RNAi enhances the survival of Leishmania.</a> Parasites and Vectors. 2 (1), 62 (2009).</li><li>Diaz-Albiter, H., Mitford, R., Genta, F. A., Sant'Anna, M. R., Dillon, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reactive+oxygen+species+scavenging+by+catalase+is+important+for+female+Lutzomyia+longipalpis+fecundity+and+mortality.">Reactive oxygen species scavenging by catalase is important for female Lutzomyia longipalpis fecundity and mortality.</a> PLoS One. 6 (3), 17486 (2011).</li><li>Barrangou, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR+provides+acquired+resistance+against+viruses+in+prokaryotes.">CRISPR provides acquired resistance against viruses in prokaryotes.</a> Science. 315 (5819), 1709-1712 (2007).</li><li>Jinek, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+programmable+dual-RNA-guided+DNA+endonuclease+in+adaptive+bacterial+immunity.">A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity.</a> Science. 337 (6096), 816-821 (2012).</li><li>Pawelczak, K. S., Gavande, N. S., VanderVere-Carozza, P. S., Turchi, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulating+DNA+Repair+Pathways+to+Improve+Precision+Genome+Engineering.">Modulating DNA Repair Pathways to Improve Precision Genome Engineering.</a> ACS Chemical Biology. 13 (2), 389-396 (2018).</li><li>Martin-Martin, I., Aryan, A., Meneses, C., Adelman, Z. N., Calvo, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimization+of+sand+fly+embryo+microinjection+for+gene+editing+by+CRISPR/Cas9.">Optimization of sand fly embryo microinjection for gene editing by CRISPR/Cas9.</a> PLoS Neglected Tropical Diseases. 12 (9), 0006769 (2018).</li><li>Meuti, M., Harrell, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparing+and+Injecting+Embryos+of+Culex+Mosquitoes+to+Generate+Null+Mutations+Using+CRISPR/Cas9.">Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations Using CRISPR/Cas9.</a> JoVE. (163), e61651 (2020).</li><li>Chaverra-Rodriguez, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+delivery+of+CRISPR-Cas9+ribonucleoprotein+into+arthropod+ovaries+for+heritable+germline+gene+editing.">Targeted delivery of CRISPR-Cas9 ribonucleoprotein into arthropod ovaries for heritable germline gene editing.</a> Nature Communications. 9 (1), 3008 (2018).</li><li>Chaverra-Rodriguez, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Germline+mutagenesis+of+Nasonia+vitripennis+through+ovarian+delivery+of+CRISPR-Cas9+ribonucleoprotein.">Germline mutagenesis of Nasonia vitripennis through ovarian delivery of CRISPR-Cas9 ribonucleoprotein.</a> Insect Molecular Biology. , (2020).</li><li>Heu, C. C., McCullough, F. M., Luan, J., Rasgon, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR-Cas9-Based+Genome+Editing+in+the+Silverleaf+Whitefly+(Bemisia+tabaci).">CRISPR-Cas9-Based Genome Editing in the Silverleaf Whitefly (Bemisia tabaci).</a> CRISPR Journal. 3 (2), 89-96 (2020).</li><li>Macias, V. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cas9-Mediated+Gene-Editing+in+the+Malaria+Mosquito+Anopheles+stephensi+by+ReMOT+Control.">Cas9-Mediated Gene-Editing in the Malaria Mosquito Anopheles stephensi by ReMOT Control.</a> Genes, Genomes and Genetics (Bethesda). 10 (4), 1353-1360 (2020).</li></ol>

We present here a recently developed embryo microinjection method for targeted mutagenesis by CRISPR/Cas9 in Phlebotomus papatasi sand flies. Embryo microinjection for insect genetic modification was developed in Drosophila in the mid-1980s21&#160;and is now routinely used in a wide variety of insects. Other methods for delivery of genetic modification materials have been developed for use in insects, such as ReMOT20,21<s...

Vector-borne diseases are a major public health threat in constant evolution. Hundreds of vector species spread across very distinct phylogenic families (e.g., mosquitoes, ticks, fleas) are responsible for the transmission of a huge number of microbial pathogens, resulting in more than 700,000 human deaths a year, according to the World Health Organization. Among vector insects, phlebotomine sand flies (Diptera, Psychodidae) constitute a vast group, with 80 proven vector species exhibiting distinct phenotypic traits and vectorial capacities found in different geographical regions. They are vectors for the protozoan parasites of the genus Leishmania, causing around 1.3 million new cases of Leishmaniases and between 20,000 and 30,000 deaths a year. Leishmaniases clinical outcomes are diverse, with symptoms ranging from self-limiting cutaneous lesions to visceral dissemination which is fatal in the absence of treatment.
Sand flies are strictly terrestrial insects. Their life cycle, relatively long compared to other Diptera, lasts up to three months, depending on different parameters such as temperature, humidity, and nutrition. It consists of one embryonic stage (6 to 11 days), four larval stages (lasting a total of 23 to 25 days) and one pupal stage (9 to 10 days) followed by metamorphosis and then adulthood. Sand flies require a humid and warm environment for rearing. Both males and females feed on sugars, obtained in the wild from flower nectars. Only females are blood-feeders, as they require proteins obtained from the blood meal for egg production1.
An important focus of research is to identify the nature of the vector/parasite interactions that lead to the development of transmissible infections. As with other vector insects, parameters intrinsic to sand flies have been shown to impact their vector competence, which is defined as their ability to carry and transmit pathogens to their hosts. For example, the expression of galectins by the Phlebotomus papatasi sand fly midgut cells, acting as receptors recognizing parasite surface components, can directly influence their vector competence for Leishmania major2,3. The insect immune response pathway, Immune Deficiency (IMD), is also crucial for the Phlebotomus papatasi sand fly vector competence for Leishmania major4. A critical role for vector insect immune response pathways in controlling their transmission of infectious pathogens has been similarly reported in Aedes aegypti mosquitos5,6,7, in the tsetse fly Glossina morsitans8, and in Anopheles gambiae mosquitoes9,10.
Studies of sand fly/Leishmania interactions have been limited by the lack of gene expression modification methods adapted for use in these insects. Only gene downregulation by small interfering RNA (siRNA) had been performed11,12,13,14 until recently. The technique, limited by the mortality associated with the microinjection of adult females, leads only to a partial loss of function, which cannot be transmitted from generation to generation.
CRISPR/Cas9 technology has revolutionized functional genomic research in non-model organisms such as sand flies. Modified from the adaptive immune system in prokaryotes for defense against bacteriophages15,16, the CRISPR Cas9 system has been rapidly adapted as a genome editing tool for superior eukaryotic organisms, including insects. The principle of CRISPR/Cas9 targeted genome-editing is based on the complementarity of a single guide RNA (sgRNA) to a specific genomic locus. The Cas9 nuclease binds to the sgRNA and creates a double-strand DNA (dsDNA) break in the genomic DNA where the sgRNA associates with its complementary sequence. The Cas9-sgRNA complex is guided to the target sequence by 17 to 20 complementary bases in the sgRNA to the chosen locus, the dsDNA break can then be repaired by two independent pathways: nonhomologous end joining (NHEJ) or homology-directed repair (HDR)17. NHEJ repair involves a simple closure of the break but frequently leads to small insertion/deletion events. DNA repair through HDR uses a donor DNA molecule sharing homology with the target DNA as a template for repair. Insects possess both machineries.
CRISPR/Cas9 technology can generate mutations in a chosen locus, through the NHEJ repair pathway; or for more complex genome editing strategies, such as knock-ins or expression reporters, through the HDR pathway with an appropriate donor template. In sand flies, null mutant alleles of the immune response factor Relish were generated through NHEJ-mediated CRISPR in Phlebotomus papatasi4. Sand fly embryos were also injected in another study with a CRISPR/Cas9 mix targeting the gene encoding Yellow. Still, no adults carrying the mutation were produced18. We describe here a detailed method of sand fly targeted mutagenesis by NHEJ-mediated CRISPR/Cas9, with a particular focus on the embryo microinjection, a critical step of the protocol.

<table>
	<tbody>
		<tr>
			<td>Black Filter Paper 4.25CM PK100</td>
			<td>VWR</td>
			<td>28342-012</td>
			<td>Cut into rectangles that are approximately 46 X 22mm. These are placed between the slide and the coverslip and act as a moist base layer for the embryos during injection.</td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>Fisher Scientific</td>
			<td>12-543A</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting Microscope</td>
			<td>Any brand</td>
			<td></td>
			<td>For aligning embryos</td>
		</tr>
		<tr>
			<td>Glass slides</td>
			<td>Fisher Scientific</td>
			<td>12-550-A3</td>
			<td>Base layer of the microinjection set up Figure 2A</td>
		</tr>
		<tr>
			<td>Insect cage</td>
			<td>custom made or several commercial options</td>
			<td></td>
			<td>polycarbonate cage for adults holding and mating Lawyer, Phillip, Mireille Killick-Kendrick, Tobin Rowland, Edgar Rowton, and Petr Volf. &#8220;Laboratory Colonization and Mass Rearing of Phlebotomine Sand Flies (Diptera, Psychodidae).&#8221; Parasite 24. Accessed August 6, 2020. https://doi.org/10.1051/parasite/2017041.</td>
		</tr>
		<tr>
			<td>Larval food</td>
			<td>custom made</td>
			<td></td>
			<td>a mix of rabbit chow and rabbit feces Lawyer, Phillip, Mireille Killick-Kendrick, Tobin Rowland, Edgar Rowton, and Petr Volf. &#8220;Laboratory Colonization and Mass Rearing of Phlebotomine Sand Flies (Diptera, Psychodidae).&#8221; Parasite 24. https://doi.org/10.1051/parasite/2017041.</td>
		</tr>
		<tr>
			<td>Microcaps 100 ml</td>
			<td>Drummond</td>
			<td>1-000-1000</td>
			<td>Used to back fill microinjection needles</td>
		</tr>
		<tr>
			<td>Mouth aspirator</td>
			<td>John W. Hock Company</td>
			<td>Model 612</td>
			<td>mouth aspirator with HEPA filter</td>
		</tr>
		<tr>
			<td>Olympus SZX12</td>
			<td>Olympus Life Sciences</td>
			<td></td>
			<td>Microinjection microscope</td>
		</tr>
		<tr>
			<td>Ovipots</td>
			<td>Nalge company</td>
			<td></td>
			<td>ovipots are made from 125-ml or 500-ml straigh-sided plolypropylene jars modified by drilling 2.5cm holes in the bottom and filled with 1cm of plaster of Paris. Lawyer, Phillip, Mireille Killick-Kendrick, Tobin Rowland, Edgar Rowton, and Petr Volf. &#8220;Laboratory Colonization and Mass Rearing of Phlebotomine Sand Flies (Diptera, Psychodidae).&#8221; Parasite 24. Accessed August 6, 2020. https://doi.org/10.1051/parasite/2017041.</td>
		</tr>
		<tr>
			<td>Paint Brush 6-0</td>
			<td>Any Art Supply Company</td>
			<td>n/a</td>
			<td>Used for aligning embryos</td>
		</tr>
		<tr>
			<td>Propionic acid</td>
			<td>Sigma-Aldrich</td>
			<td>402907</td>
			<td>antifungal agent</td>
		</tr>
		<tr>
			<td>Standard Glass Capillaries</td>
			<td>World Precision Instruments</td>
			<td>1B100-3</td>
			<td>Used for making microinjection needles</td>
		</tr>
		<tr>
			<td>Trio-MPC100 Controller and MP845 Manipulator</td>
			<td>Sutter Instruments</td>
			<td></td>
			<td>Microinjection Controller and Micromanipulator</td>
		</tr>
	</tbody>
</table>

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.

The CRISPR/Cas9 microinjection protocol described here to generate sand fly mutants was established in a previous publication4. This approach produced highly efficient mutagenesis, as 11 out of 540 individuals survived the procedure, of which 9 were mutant. When designing guides for CRISPR/Cas9 mutation, a critical first step is to sequence the region around the area to be targeted. The template for sequencing should be from the strain that is going to be used as a source of embryos for injection....

Watch this Scientific Journal Video about Sand Fly Phlebotomus papatasi Embryo Microinjection for CRISPR/Cas9 Mutagenesis at JoVE.com

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 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 ...

Targeted mutagenesis has been adapted to sandflies only recently and is a crucial technique for understanding the role and function of genes of interest in these vector insects. Embryo microinjection is a very important step in insect genome editing, but it needs to be adapted to species that you are working with, in our case, sandflies. Compared to the other insect species, sand fly embryos are very small.Take a long time to develop and are particularly sensitive to the level of humidity. Five days before the injection, blood feed female sandflies as for routine colony maintenance. One day post feeding, capture the blood-fed females in plaster pods with a side port in groups of 100 to 150 and feed the flies with a 30%sucrose solution.On day five post-blood feeding, place a white piece of moist filter paper into the egg-laying chamber. And use a mouth aspirator to collect and transfer about 10 females from the plaster pod to the egg-laying chamber. After 30 to 60 minutes, remove the Petri dish and filter paper from the egging chamber.Taking care of that the filter paper remains moist and continuing to collect new groups of females and embryos throughout this period. When all of the embryos have been collected, use a stereo microscope and a very fine paint brush to carefully transfer about 50 embryos onto individual pieces of moist black filter paper, place onto microscope slides, topped with a single cover slip per slide. Add enough water to each filter paper to keep the embryos moist without floating or being sucked under the cover slips and line the embryos against the cover slips.Two to three hours after the start of the egg collection, place a slide under a dissecting microscope and use the paintbrush to carefully add water to the back end of the filter paper. Back load 0.5 to one microliters of load injection mix into a micro injection needle and mount the needle into a pneumatic injection controller, set to 30 pounds per square inch of pressure under the microscope. Depress the trigger to expel any air from the needle tip.When the air has been released, slowly increase the hold pressure. When the injection material starts to flow, decrease the hold pressure until it is just below the point of the injection mix flowing from the needle and insert the needle into the side of the first embryo. Using the cover slip behind the embryo as a back stop, deliver a small amount of injection mix into the embryo before gently removing the needle.Immediately after removing the needle, press the injector to remove any back-filled material from the needle tip and proceed to the next embryo. When all of the embryos have been injected, count the number of injected embryos to keep a running tally and add water to the filter paper so that the cover slip floats. Then use a probe to hold the filter paper in place while pulling the cover slip away from the embryos.And he was filter paper to blot away the excess water from the slide. Transfer the filter paper onto a stack of moist filter paper in a new Petri dish and manually transfer each embryo onto moistened, small sized plaster pods without crowding. Then cover the pods with a screen and store the pods in a 26 degree incubator at a 70%humidity.As with most successful microinjection protocols, a good microinjection needle suited to the embryos being injected is important. Good sharp needles can easily penetrate the embryo without allowing the material to escape post-injection. The pulled needles should not have a taper that is too long.Otherwise, the lumen of the needle becomes very narrow for a major portion of the taper, making it difficult to get the injection pressure high enough to force the material through the needle. Post-injection reared embryos that are healthy should be retained, while damaged fungus contaminated, desiccated or deformed embryos should be discarded. Potential mutant alleles can be identified in post-injection reared individuals by PCR analysis of the region surrounding the expected cutting sites.After mutant line establishment using the injected embryos, genotyping PCR allows the identification of homozygous mutant sibling crosses. It's essential to have a good needle to be gentle and manipulating and injecting the embryos. And to make sure that the embryos are hydrated throughout the entire procedure.Genome editing in some flies is just getting started. This technique is really a first step that should be followed by more complex genome modification strategies.

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Research

JoVE Journal

Genetics

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 ...

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene function. Gene knockout has been used to study these gene functions in vivo. The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system is a powerful tool used to edit genomic DNA sequences to alter gene function. While the traditional approach has been to introduce CRISPR/Cas9 mRNA into the single cell embryos through microinjection, this can be a slow and inefficient process in catfish. Here, a detailed protocol for microinjection of channel catfish embryos with CRISPR/Cas9 protein is described. Briefly, eggs and sperm were collected and then artificial fertilization performed. Fertilized eggs were transferred to a Petri dish containing Holtfreter's solution. Injection volume was calibrated and then guide RNAs/Cas9 targeting the toll/interleukin 1 receptor domain-containing adapter molecule (TICAM 1) gene and rhamnose binding lectin (RBL) gene were microinjected into the yolk of one-cell embryos. The gene knockout was successful as indels were confirmed by DNA sequencing. The predicted protein sequence alterations due to these mutations included frameshift and truncated protein due to premature stop codons.

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

A simple and efficient microinjection protocol for gene editing in channel catfish embryos using the CRISPR/Cas9 system is presented. In this protocol, guide RNAs and Cas9 protein were microinjected into the yolk of one-cell embryos. This protocol has been validated by knocking out two channel catfish immune-related genes.

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene ...

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration of transgenes. However, due to the low efficiency of homologous recombination (HR) and various indel mutations of non-homologous end joining (NHEJ)-based strategies in non-dividing cells, in vivo genome editing remains a great challenge. Here, we describe a homology-mediated end joining (HMEJ)-based CRISPR/Cas9 system for efficient in vivo precise targeted integration. In this system, the targeted genome and the donor vector containing homology arms (~800 bp) flanked by single guide RNA (sgRNA) target sequences are cleaved by CRISPR/Cas9. This HMEJ-based strategy achieves efficient transgene integration in mouse zygotes, as well as in hepatocytes in vivo. Moreover, a HMEJ-based strategy offers an efficient approach for correction of fumarylacetoacetate hydrolase (Fah) mutation in the hepatocytes and rescues Fah-deficiency induced liver failure mice. Taken together, focusing on targeted integration, this HMEJ-based strategy provides a promising tool for a variety of applications, including generation of genetically modified animal models and targeted gene therapies.

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system provides a promising tool for genetic engineering, and opens up the possibility of targeted integration of transgenes. We describe a homology-mediated end joining (HMEJ)-based strategy for efficient DNA targeted integration in vivo and targeted gene therapies using CRISPR/Cas9.

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration ...

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex determination, venom synthesis, and host-symbiont interactions, among others. A major limitation of working with this organism is the lack of effective protocols to perform directed genome modifications. An important part of genome modification is delivery of editing reagents, including CRISPR/Cas9 molecules, into embryos through microinjection. While microinjection is well established in many model organisms, this technique is particularly challenging to perform in N. vitripennis primarily due to its small embryo size, and the fact that embryonic development occurs entirely within a parasitized blowfly pupa. The following procedure overcomes these significant challenges while demonstrating a streamlined, visual procedure for effectively removing wasp embryos from parasitized host pupae, microinjecting them, and carefully transplanting them back into the host for continuation and completion of development. This protocol will strongly enhance the capability of research groups to perform advanced genome modifications in this organism.

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Microinjection of Nasonia vitripennis embryos is an essential method for generating heritable genome modifications. Described here is a detailed procedure for microinjection and transplantation of Nasonia vitripennis embryos, which will greatly facilitate future genome manipulation in this organism.

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex ...

Endogenous Protein Tagging in Human Induced Pluripotent Stem Cells Using CRISPR/Cas9

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or C-terminal fluorescent tags. The prokaryotic CRISPR/Cas9 system (clustered regularly interspaced short palindromic repeats/CRISPR-associated 9) may be used to introduce large exogenous sequences into genomic loci via homology directed repair (HDR). To achieve the desired knock-in, this protocol employs the ribonucleoprotein (RNP)-based approach where wild type Streptococcus pyogenes Cas9 protein, synthetic 2-part guide RNA (gRNA), and a donor template plasmid are delivered to the cells via electroporation. Putatively edited cells expressing the fluorescently tagged proteins are enriched by fluorescence activated cell sorting (FACS). Clonal lines are then generated and can be analyzed for precise editing outcomes. By introducing the fluorescent tag at the genomic locus of the gene of interest, the resulting subcellular localization and dynamics of the fusion protein can be studied under endogenous regulatory control, a key improvement over conventional overexpression systems. The use of hiPSCs as a model system for gene tagging provides the opportunity to study the tagged proteins in diploid, nontransformed cells. Since hiPSCs can be differentiated into multiple cell types, this approach provides the opportunity to create and study tagged proteins in a variety of isogenic cellular contexts.

Described here is a protocol for tagging endogenously expressed proteins with fluorescent tags in human induced pluripotent stem cells using CRISPR/Cas9. Putatively edited cells are enriched by fluorescence activated cell sorting and clonal cell lines are generated.

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or ...

CRISPR/Cas9 Ribonucleoprotein-mediated Precise Gene Editing by Tube Electroporation

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of CRISPR/Cas9 elements into the target cells by transfection is a prerequisite for efficient gene editing. This protocol demonstrates that tube electroporation (TE) machine-mediated delivery of CRISPR/Cas9 ribonucleoprotein (RNP), along with single-stranded oligodeoxynucleotide (ssODN) donor templates to different types of mammalian cells, leads to robust precise gene editing events. First, TE was applied to deliver CRISPR/Cas9 RNP and ssODNs to induce disease-causing mutations in the interleukin 2 receptor subunit gamma (IL2RG) gene and sepiapterin reductase (SPR) gene in rabbit fibroblast cells. Precise mutation rates of 3.57%-20% were achieved as determined by bacterial TA cloning sequencing. The same strategy was then used in human iPSCs on several clinically relevant genes including epidermal growth factor receptor (EGFR), myosin binding protein C, cardiac (Mybpc3), and hemoglobin subunit beta (HBB). Consistently, highly precise mutation rates were achieved (11.65%-37.92%) as determined by deep sequencing (DeepSeq). The present work demonstrates that tube electroporation of CRISPR/Cas9 RNP represents an efficient transfection protocol for gene editing in mammalian cells.

Presented here is a protocol for efficient CRISPR/Cas9 ribonucleoprotein-mediated gene editing in mammalian cells using tube electroporation.

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of ...

Generation of Defined Genomic Modifications Using CRISPR-CAS9 in Human Pluripotent Stem Cells

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise heterozygous genetic modifications is challenging due to CRISPR-CAS9 mediated indel formation in the second allele. Here, we demonstrate a protocol to help overcome this difficulty by using two repair templates in which only one expresses the desired sequence change, while both templates contain silent mutations to prevent re-cutting and indel formation. This methodology is most advantageous for gene editing coding regions of DNA to generate isogenic control and mutant human stem cell lines for studying human disease and biology. In addition, optimization of transfection and screening methodologies have been performed to reduce labor and cost of a gene editing experiment. Overall, this protocol is widely applicable to many genome editing projects utilizing the human pluripotent stem cell model.

This protocol provides a method to facilitate the generation of defined heterozygous or homozygous nucleotide changes using CRISPR-CAS9 in human pluripotent stem cells.

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise ...

CRISPR-Cas9-Mediated Genome Editing in the Filamentous Ascomycete Huntiella omanensis

The CRISPR-Cas9 genome editing system is a molecular tool that can be used to introduce precise changes into the genomes of model and non-model species alike. This technology can be used for a variety of genome editing approaches, from gene knockouts and knockins to more specific changes like the introduction of a few nucleotides at a targeted location. Genome editing can be used for a multitude of applications, including the partial functional characterization of genes, the production of transgenic organisms and the development of diagnostic tools. Compared to previously available gene editing strategies, the CRISPR-Cas9 system has been shown to be easy to establish in new species and boasts high efficiency and specificity. The primary reason for this is that the editing tool uses an RNA molecule to target the gene or sequence of interest, making target molecule design straightforward, given that standard base pairing rules can be exploited. Similar to other genome editing systems, CRISPR-Cas9-based methods also require efficient and effective transformation protocols as well as access to good quality sequence data for the design of the targeting RNA and DNA molecules. Since the introduction of this system in 2013, it has been used to genetically engineer a variety of model species, including Saccharomyces cerevisiae, Arabidopsis thaliana, Drosophila melanogaster and Mus musculus. Subsequently, researchers working on non-model species have taken advantage of the system and used it for the study of genes involved in processes as diverse as secondary metabolism in fungi, nematode growth and disease resistance in plants, among many others. This protocol detailed below describes the use of the CRISPR-Cas9 genome editing protocol for the truncation of a gene involved in the sexual cycle of Huntiella omanensis, a filamentous ascomycete fungus belonging to the Ceratocystidaceae family.

CRISPR-Cas9-Mediated Genome Editing in the Filamentous Ascomycete Huntiella omanensis

The CRISPR-Cas9 genome editing system is an easy-to-use genome editor that has been used in model and non-model species. Here we present a protein-based version of this system that was used to introduce a premature stop codon into a mating gene of a non-model filamentous ascomycete fungus.

The CRISPR-Cas9 genome editing system is a molecular tool that can be used to introduce precise changes into the genomes of model and non-model species ...

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, ...

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

The unique traits of eusocial insects, such as social behavior and reproductive division of labor, are controlled by their genetic system. To address how genes regulate social traits, we have developed mutant ants via delivery of CRISPR complex into young embryos during their syncytial stage. Here, we provide a protocol of CRISPR-mediated mutagenesis in Harpegnathos saltator, a ponerine ant species that displays striking phenotypic plasticity. H. saltator ants are readily reared in a laboratory setting. Embryos are collected for microinjection with Cas9 proteins and in vitro synthesized small guide RNAs (sgRNAs) using home-made quartz needles. Post-injection embryos are reared outside the colony. Following emergence of the first larva, all embryos and larvae are transported to a nest box with a few nursing workers for further development. This protocol is suitable for inducing mutagenesis for analysis of caste-specific physiology and social behavior in ants, but may also be applied to a broader spectrum of hymenopterans and other insects.

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

Many characteristics of insect eusociality rely on within-colony communication and division of labor. Genetic manipulation of key regulatory genes in ant embryos via microinjection and CRISPR-mediated mutagenesis provides insights into the nature of altruistic behavior in eusocial insects.

The unique traits of eusocial insects, such as social behavior and reproductive division of labor, are controlled by their genetic system. To address how ...