All figures were created with Biorender.com under a license to Texas A&#38;M University. This work was supported by funds from the National Institute of Allergy and Infectious Disease (AI137112 and AI115138 to Z.N.A.), Texas A&#38;M AgriLife Research under the Insect Vectored Disease Grant Program, and the USDA National Institute of Food and Agriculture, Hatch project 1018401.

1. Scanning for single nucleotide polymorphisms (SNPs), HRMA primer design, and primer validation
<ol>
	<li>SNP identification in wild-type laboratory colony mosquitoes
	<ol>
		<li>Select the target exon to disrupt proper polypeptide translation. 
		NOTE: The target should be close to the start codon or amongst key residues required for protein function. The shorter the exon (e.g., &#8804;200 bases), the more difficult it is to target and analyze. Avoid editing close to the boundaries of an exon, as this forces one of the HRMA primers to either cross an intron or be in an intron. This is undesirable because SNP rates tend to be much greater in those regions.</li>
		<li>Primer design
		<ol>
			<li>Go to the NCBI Blast - Primer Blast website15, copy and paste the chosen exon in the box at the top of the page | choose the PCR product size (ensure it encompasses most of the exon) | choose the organism.</li>
			<li>Click on Advanced parameters | Opt (for PCR Product Tm) and add 60 (for an optimal temperature of 60 &#176;C) | Get Primers. Keep the other parameters as default.</li>
		</ol>
		</li>
		<li>Obtain genomic DNA (gDNA) from the wild-type laboratory colony. Set aside 10 mosquitoes, anesthetize them with CO2,&#160;and place them in a Petri dish on ice to keep them inactive. Set up 10 tubes containing 0.5 &#181;L of the reagent for release of DNA from tissue and 20 &#181;L of dilution buffer, both provided in the DNA release kit suggested in the Table of Materials.</li>
		<li>Remove one leg from a single mosquito using forceps and place it in a corresponding tube of a diluted solution of the DNA-release reagent (from step 1.1.3), completely submerging the leg in the solution. Repeat this step with the remaining mosquitoes, wiping the forceps with 70% ethanol before proceeding to the next one.</li>
		<li>Incubate the leg-containing solution at room temperature for 2-5 min, then at 98 &#176;C for 2 min, and allow it to cool down while setting up the PCR reaction. 
		NOTE: Plates containing the released gDNA can be stored at -20 &#176;C, and the protocol can be paused at this point.</li>
		<li>Prepare 10 tubes containing 10 &#181;L of 2x PCR Master Mix, primers to a 0.5 &#181;M final concentration, and molecular grade water to a final volume of 20 &#181;L, and transfer 1 &#181;L of the diluted sample to each tube. Perform PCR following these cycling parameters: 98 &#176;C 5 min, 40 cycles of 98 &#176;C for 5 s, 60 &#176;C 30 s, 72 &#176;C 20 s per kb; final extension of 72 &#176;C for 1 min.</li>
		<li>Purify the PCR products with an enzyme to degrade the residual PCR primers and dephosphorylate excess dNTPs or any column clean-up kit. Proceed with sequencing the samples.</li>
		<li>Analyze each electropherogram for the presence of double peaks/ambiguous bases, and adjust base calls manually in each sequence using the appropriate degenerate base code. 
		NOTE: This step must be performed before multiple sequence alignment as it is common for the base-calling software to select the more prominent peak as the &#34;true&#34; base call, giving the false impression of an absence of SNPs.</li>
		<li>Perform a multiple sequence alignment using the alignment software, SeqMan Pro, listed in the Table of Materials or other open-source alignment software, such as ClustalW16 or T-Coffee17.
		<ol>
			<li>Open the alignment software | click on Add Sequences | select the desired sequences and click on Add | once all sequences are chosen, click on Done.</li>
			<li>Click on Assemble to perform the alignment. To open the alignment, click on Contig 1 and analyze the alignment and identify the SNPs (Figure 1). 
			NOTE: As an alternative to steps 1.1.3-1.1.6, PCR can be performed from isolated genomic DNA obtained from bulk samples (derived from &#62;10 individuals). Sequenced amplicons can be analyzed directly for the presence of SNPs appearing as double peaks in the electropherogram, though rare SNPs will be more difficult to detect.</li>
		</ol>
		</li>
		<li>Design 3-5 single guide RNAs (sgRNAs), avoiding regions containing any SNP identified above, following the protocol described in18.</li>
	</ol>
	</li>
	<li>HRMA primer design
	<ol>
		<li>Test the exon sequence for the possible formation of secondary structures during PCR using mFold19.
		<ol>
			<li>Go to the UNAFold Web Server | click on mFold | on the dropdown menu, click on Applications | DNA Folding Form.</li>
			<li>Enter the sequence name in the box and paste the exon.</li>
			<li>Change the folding temperature to 60 &#176;C; on the Ionic conditions, change [Mg++] to 1.5 and on Units switch to mM; click on Fold DNA.</li>
			<li>On Output, below Structure 1, click on pdf to open the Circular Structure Plots.&#160;</li>
		</ol>
		</li>
		<li>Go to NCBI Blast - Primer Blast15 for primer design.</li>
		<li>Copy and paste the selected exon sequence determined by sequencing in step 1.1 (the sequence that contains the lowest number of SNPs or no SNPs) in the box at the top of the page.</li>
		<li>Use the symbol &#60; &#62; to mark and exclude sequences that contain SNPs, the target site, and regions with possible formation of secondary structure.</li>
		<li>Select the PCR product size to be between 80 and 150 bp and choose the organism. 
		NOTE: Larger fragment sizes can be successfully used (~300 bp). However, longer amplicons may decrease sensitivity between sequences differing in one or just a few base pairs.</li>
		<li>Click on Get Primers. Select 2&#8211;3 pairs of primers to be tested (ideally, primer sites are &#8805;20&#8211;50 bp away from any CRISPR target sites).</li>
	</ol>
	</li>
	<li>Primer validation
	<ol>
		<li>Perform a gradient PCR using gDNA from a single individual.
		<ol>
			<li>Prepare a master mix and remove one sample for the non-template control (NTC) in a separate tube. Add the template to the remaining master mix and aliquot into a 96-well plate.</li>
			<li>Follow the cycling parameters: 98 &#176;C 30 s, 34 cycles of 98 &#176;C for 10 s, 55&#8211;65 &#176;C 30 s, 72 &#176;C 15 s; final extension of 72 &#176;C for 10 min.</li>
			<li>Generate thermal melt profiles following the parameters: denaturation step 95 &#176;C for 1 min, annealing 60 &#176;C for 1 min, melt curve detection between 75 &#176;C and 95 &#176;C in 0.2 &#176;C increments, with a hold time of 10 s at each temperature. 
			NOTE: Only annealing temperatures with a single thermal melt profile should be used.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Proceed to generate the mutant lines with embryo injections as described in12.</li>
</ol>
2. Preparation of genomic DNA from mosquito legs
<ol>
	<li>Separate the G1 mosquitoes by sex at the pupal stage so that they do not mate before genotyping, and make sure that age-matched, wild-type control mosquitoes will be available to be used for references and backcrosses. Sex-separate them likewise.</li>
	<li>Gather the materials needed (Figure 2A) and prepare a 96 well PCR plate with 0.5 &#181;L of the DNA-release reagent and 20 &#181;L of dilution buffer (from the gDNA release kit) per individual to be genotyped (Figure 2B) and leave it on ice. Reserve two reactions for NTC.</li>
	<li>Label a tray for mosquito vials (Drosophila vials) so each well on the 96-plate corresponds to the respective mosquito vial in the tray.</li>
	<li>Anesthetize the G1 mosquitos with CO2 and place them in a glass Petri dish to keep them sedated (Figure 2C,D). Anesthetize and place 8 wild-type mosquitoes in a second Petri dish.</li>
	<li>Wipe a pair of tweezers with 70% ethanol and remove one of the mosquito hind legs (Figure 2E,F).</li>
	<li>Submerge the leg in the DNA-release reagent solution, place the mosquito in the corresponding vial, and close it with a sponge (Figure 2G,H).</li>
	<li>Wipe the tweezers again with 70% ethanol and proceed with removing the leg from the next mosquito. Repeat steps 2.5-2.7 until the 96-well plate is completed. 
	NOTE: It is important to wipe the tweezers with 70% ethanol. This minimizes DNA cross-contamination.</li>
	<li>Seal the plate with an optical PCR plate seal (Figure 2I) and incubate the 96-well plate containing the legs at room temperature (RT) for 2-5 min and then at 98 &#176;C for 2 min. Allow the plate to cool down to RT while preparing the PCR mix. 
	NOTE: The entire process typically takes 3-4 h. If the mosquitoes must be kept in the vials for more time than the expected duration (especially &#8805;1 day), place a small piece of raisin (or source of sugar and water) with each mosquito to ensure the survival of the mosquitoes during extended incubations.</li>
</ol>
3. HRMA
<ol>
	<li>Perform PCR.
	<ol>
		<li>Prepare a master mix containing the following components per each reaction: 10 &#181;L of 2x buffer (from the gDNA release kit), 0.5 &#181;M of each primer, 1 &#181;L of EvaGreen dye, 0.4 &#181;L of polymerase (from gDNA release kit), and complete to 19 &#181;L with molecular-grade water.</li>
		<li>Using a multichannel pipet, transfer 19 &#181;L of the master mix into each well. Transfer 1 &#181;L of the DNA release solution containing mosquito DNA prepared in section 2 to the plate (Figure 3A). Seal the plate with an optical PCR plate seal.</li>
		<li>Perform the PCR following the cycling parameters: 98 &#176;C for 5 min, 39 cycles of 98 &#176;C for 10 s, the chosen annealing temperature (72 &#176;C) for 30 s; final extension 72 &#176;C for 2 min.</li>
	</ol>
	</li>
	<li>Generate thermal melt profiles following the parameters: denaturation step 95 &#176;C for 1 min, annealing 60 &#176;C for 1 min, melt curve detection between 75 &#176;C and 95 &#176;C in 0.2 &#176;C increments, with a hold time of 10 s at each temperature. See the Supplemental Material and Supplemental Figure S1-S5 for a detailed description of the software setup for HRMA run using the CFX96 Real-Time System.</li>
	<li>Examine the melt profiles (Figure 3B). Assign wild-type control to the reference cluster. 
	NOTE: The software (see the Table of Materials) automatically normalizes the data and designates clusters with colors for different melt curves.</li>
	<li>Mark the different clusters with corresponding colors on the 96-well template (Figure 3C).</li>
	<li>Select the individuals with curves of interest, remove them from the tubes, and backcross (Figure 3D). Blood-feed the mated females and collect G2 eggs.</li>
</ol>
4. Sequence verification by Sanger sequencing
<ol>
	<li>Purify the PCR product from the wells with selected mosquitoes (from the plate prepared in section 3.1), using an enzyme to degrade the residual PCR primers, and dephosphorylate excess dNTPs. Alternatively, use any column clean-up kit and proceed with sequencing the samples. 
	NOTE: Direct sequencing may be more challenging when the PCR product size is small (&#8804;200 bp). In such cases, design primers to amplify larger fragments encompassing the target site (&#8805;250 bp) and amplify by PCR using the DNA in the 96-well plate (step 2.2).</li>
	<li>Analyze and identify the indels using trace viewer software (Figure 4). Alternatively, Poly Peak Parser software20 can help detect the mutation in heterozygous individuals.
	<ol>
		<li>Go to the Poly Peak Parser website, select the sequence from the mutant individuals on the Browse menu, and copy and paste the reference sequence on the box. 
		NOTE: The alignment between the alternate allele and the reference will appear automatically on the right side of the screen.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>WHO. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dengue+Guidelines+for+diagnosis,+treatment,+prevention+and+control.">Dengue Guidelines for diagnosis, treatment, prevention and control.</a> WHO. , 160 (2009).</li><li>WHO. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zika+epidemiology+update.">Zika epidemiology update.</a> WHO. , 1-13 (2019).</li><li>WHO. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Guidelines+for+prevention+and+control+of+Chikungunya+fever.">Guidelines for prevention and control of Chikungunya fever.</a> WHO. , (2009).</li><li>World malaria report 2020: 20 years of global progress and challenges. World Health Organization Available from: <a target="_blank" href="https://www.who.int/publications/i/item/9789240015791">https://www.who.int/publications/i/item/9789240015791</a> (2020)</li><li>Qiu, P., 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=Mutation+detection+using+Surveyor+nuclease.">Mutation detection using Surveyor nuclease.</a> Biotechniques. 36 (4), 702-707 (2004).</li><li>Babon, J. J., McKenzie, M., Cotton, R. 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+use+of+resolvases+T4+endonuclease+VII+and+T7+endonuclease+I+in+mutation+detection.">The use of resolvases T4 endonuclease VII and T7 endonuclease I in mutation detection.</a> Molecular Biotechnology. 23 (1), 73-81 (2003).</li><li>Erali, M., Voelkerding, K. V., Wittwer, C. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High+resolution+melting+applications+for+clinical+laboratory+medicine.">High resolution melting applications for clinical laboratory medicine.</a> Experimental and Molecular Pathology. 85 (1), 50-58 (2008).</li><li>Reed, G. H., Wittwer, C. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sensitivity+and+specificity+of+single-nucleotide+polymorphism+scanning+by+high-resolution+melting+analysis.">Sensitivity and specificity of single-nucleotide polymorphism scanning by high-resolution melting analysis.</a> Clinical Chemistry. 50 (10), 1748-1754 (2004).</li><li>Parant, J. M., George, S. A., Pryor, R., Wittwer, C. T., Yost, H. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+rapid+and+efficient+method+of+genotyping+zebrafish+mutants.">A rapid and efficient method of genotyping zebrafish mutants.</a> Developmental Dynamics. 238 (12), 3168-3174 (2009).</li><li>Tong, S. Y., Giffard, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microbiological+applications+of+high-resolution+melting+analysis.">Microbiological applications of high-resolution melting analysis.</a> Journal of Clinical Microbiology. 50 (11), 3418-3421 (2012).</li><li>Simko, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-resolution+DNA+melting+analysis+in+plant+research.">High-resolution DNA melting analysis in plant research.</a> Trends in Plant Science. 21 (6), 528-537 (2016).</li><li>Basu, 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=Silencing+of+end-joining+repair+for+efficient+site-specific+gene+insertion+after+TALEN/CRISPR+mutagenesis+in+Aedes+aegypti.">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 of the United States of America. 112 (13), 4038-4043 (2015).</li><li>O'Leary, S., Adelman, Z. N. <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+knockout+of+female-biased+genes+AeAct-4+or+myo-fem+in+Ae.+aegypti+results+in+a+flightless+phenotype+in+female,+but+not+male+mosquitoes.">CRISPR/Cas9 knockout of female-biased genes AeAct-4 or myo-fem in Ae. aegypti results in a flightless phenotype in female, but not male mosquitoes.</a> PLoS Neglected Tropical Diseases. 14 (12), 0008971 (2020).</li><li>Kojin, B. B., 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=Aedes+aegypti+SGS1+is+critical+for+Plasmodium+gallinaceum+infection+of+both+the+mosquito+midgut+and+salivary+glands.">Aedes aegypti SGS1 is critical for Plasmodium gallinaceum infection of both the mosquito midgut and salivary glands.</a> Malaria Journal. 20 (1), 11 (2021).</li><li>Ye, J., 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=Primer-BLAST:+a+tool+to+design+target-specific+primers+for+polymerase+chain+reaction.">Primer-BLAST: a tool to design target-specific primers for polymerase chain reaction.</a> BMC Bioinformatics. 13, 134 (2012).</li><li>Larkin, M. A., 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=Clustal+W+and+Clustal+X+version+2.0.">Clustal W and Clustal X version 2.0.</a> Bioinformatics. 23 (21), 2947-2948 (2007).</li><li>Notredame, C., Higgins, D. G., Heringa, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=T-Coffee:+A+novel+method+for+fast+and+accurate+multiple+sequence+alignment.">T-Coffee: A novel method for fast and accurate multiple sequence alignment.</a> Journal of Molecular Biology. 302 (1), 205-217 (2000).</li><li>Bassett, A. R., Tibbit, C., Ponting, C. P., Liu, 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=Highly+efficient+targeted+mutagenesis+of+Drosophila+with+the+CRISPR/Cas9+system.">Highly efficient targeted mutagenesis of Drosophila with the CRISPR/Cas9 system.</a> Cell Reports. 6 (6), 1178-1179 (2014).</li><li>Zuker, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mfold+web+server+for+nucleic+acid+folding+and+hybridization+prediction.">Mfold web server for nucleic acid folding and hybridization prediction.</a> Nucleic Acids Research. 31 (13), 3406-3415 (2003).</li><li>Hill, J. T., 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=Poly+peak+parser:+Method+and+software+for+identification+of+unknown+indels+using+sanger+sequencing+of+polymerase+chain+reaction+products.">Poly peak parser: Method and software for identification of unknown indels using sanger sequencing of polymerase chain reaction products.</a> Developmental Dynamics. 243 (12), 1632-1636 (2014).</li><li>Tsujimoto, H., Anderson, M. A. E., Myles, K. M., Adelman, Z. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+candidate+iron+transporters+from+the+ZIP/ZnT+gene+families+in+the+mosquito+Aedes+aegypti.">Identification of candidate iron transporters from the ZIP/ZnT gene families in the mosquito Aedes aegypti.</a> Frontiers in Physiology. 9, 380 (2018).</li><li>Slomka, M., Sobalska-Kwapis, M., Wachulec, M., Bartosz, G., Strapagiel, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High+resolution+melting+(HRM)+for+high-throughput+genotyping-limitations+and+caveats+in+practical+case+studies.">High resolution melting (HRM) for high-throughput genotyping-limitations and caveats in practical case studies.</a> International Journal of Molecular Sciences. 18 (11), 2316 (2017).</li><li>Vouillot, L., Thelie, A., Pollet, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+T7E1+and+surveyor+mismatch+cleavage+assays+to+detect+mutations+triggered+by+engineered+nucleases.">Comparison of T7E1 and surveyor mismatch cleavage assays to detect mutations triggered by engineered nucleases.</a> G3. 5 (3), 407-415 (2015).</li></ol>

The authors have no conflicts of interest to declare.

High-resolution melt analysis offers a simple and fast solution for the identification of indels generated by CRISPR/Cas9 technology in the vector mosquito Ae. aegypti. It provides flexibility, enabling the genotyping of mosquitoes mutated for a wide range of genes from flight muscle to iron metabolism and more13,14. HRMA can be performed in just a few hours from sample collection to the final analyses. Additional time is required for primer design and w...

As vectors of pathogens such as dengue1, Zika2, and chikungunya3 viruses, as well as malarial parasites4, mosquitoes represent a significant public health threat to humans. For all these diseases, there is a substantial focus of transmission intervention on the control of mosquito vectors. Study of the genes important in, for example, permissiveness to pathogens, mosquito fitness, survivorship, reproduction, and resistance to insecticides is key for developing novel mosquito control strategies. For such purposes, genome editing in mosquitoes is becoming a common practice, especially with the development of technologies such as HEs, ZFNs, TALENs, and most recently, CRISPR with Cas9. The establishment of gene-edited strains typically involves backcrossing individuals carrying the desired mutations for a few generations to minimize off-target and founder (bottleneck) effects, followed by crossing heterozygous individuals to generate homozygous or trans-heterozygous lines. In the absence of a dominant marker, molecular genotyping is necessary in this process because, in many cases, no clear phenotypic traits can be detected for heterozygous mutants.
Although sequencing is the gold standard for genotypic characterization, performing this across hundreds, or possibly thousands of individuals, poses significant costs, labor, and time required to obtain results, which is especially critical for organisms with short lifespans such as mosquitoes. Commonly used alternatives are Surveyor nuclease assay5 (SNA), T7E1 assay6, and high resolution melt analysis (HRMA, reviewed in7). Both SNA and T7E1 use endonucleases that cleave only mismatched bases. When a mutated region of the heterozygous mutant genome is amplified, DNA fragments from mutant and wild-type alleles are annealed to make mismatched double-stranded DNA (dsDNA). SNA detects the presence of mismatches via digestion with a mismatch-specific endonuclease and simple agarose gel electrophoresis. Alternatively, HRMA uses the thermodynamic properties of dsDNA detected by dsDNA-binding fluorescent dyes, with the disassociation temperature of the dye varying based on the presence and type of mutation. HRMA has been used for the detection of single-nucleotide polymorphisms (SNPs)8, mutant genotyping of zebra fish9, microbiological applications10, and plant genetic research11, among others.
This paper describes HRMA, a simple method of molecular genotyping for mutant mosquitoes generated by CRISPR/Cas9 technology. The advantages of HRMA over alternative techniques include 1) flexibility, as it has been proven useful for various genes, a wide range of indel sizes, as well as the distinction between different indel sizes and heterozygous, homozygous, and trans-heterozygous differentiation12,13,14, 2) cost, as it is based on commonly used PCR reagents, and 3) time-saving, as it can be performed in just a few hours. In addition, the protocol uses a small body part (a leg) as a source of DNA, allowing the mosquito to survive the genotyping process, permitting the establishment and maintenance of mutant lines.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>70% Ethanol</td>
			<td></td>
			<td></td>
			<td>70% ethanol solution in water</td>
		</tr>
		<tr>
			<td>96-well PCR and Real-time PCR plates</td>
			<td>VWR</td>
			<td>82006-636</td>
			<td>For obtaining genomic DNA (from the mosquito leg)</td>
		</tr>
		<tr>
			<td>96-well plate templates</td>
			<td></td>
			<td></td>
			<td>House-made printed, for genotype recording</td>
		</tr>
		<tr>
			<td>Bio Rad CFX96</td>
			<td>Bio Rad</td>
			<td></td>
			<td>PCR machine with gradient and HRMA capabilities</td>
		</tr>
		<tr>
			<td>Diversified Biotech reagent reservoirs</td>
			<td>VWR</td>
			<td>490006-896</td>
			<td></td>
		</tr>
		<tr>
			<td>Exo-CIP Rapid PCR Cleanup Kit</td>
			<td>New England Biolabs</td>
			<td>E1050S</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Petri Dish</td>
			<td>VWR</td>
			<td>89001-246</td>
			<td>150 mm x 20 mm</td>
		</tr>
		<tr>
			<td>Hard-shell thin-wall 96-well skirted PCR plates</td>
			<td>Bio-rad</td>
			<td>HSP9665</td>
			<td>For HRMA</td>
		</tr>
		<tr>
			<td>Multi-channel pipettor (P10)</td>
			<td>Integra Biosciences</td>
			<td>4721</td>
			<td></td>
		</tr>
		<tr>
			<td>Multi-channel pipettor (P300)</td>
			<td>Integra Biosciences</td>
			<td>4723</td>
			<td></td>
		</tr>
		<tr>
			<td>Nunc Polyolefin Acrylate Sealing tape, Thermo Scientific</td>
			<td>VWR</td>
			<td>37000-548</td>
			<td>To use with the 96-well&#160; PCR plates for obtaining genomic DNA</td>
		</tr>
		<tr>
			<td>Optical sealing tape</td>
			<td>Bio-rad</td>
			<td>2239444</td>
			<td>To use with the 96-well skirted PCR plates for HRMA</td>
		</tr>
		<tr>
			<td>Phire Animal tissue direct PCR Kit (without sampling tools)</td>
			<td>Thermo Fisher</td>
			<td>F140WH</td>
			<td>For obtaining genomic DNA and performing PCR</td>
		</tr>
		<tr>
			<td>Plastic Fly Vial Dividers</td>
			<td>Genesee</td>
			<td>59-128W</td>
			<td></td>
		</tr>
		<tr>
			<td>Precision Melt Analysis Software</td>
			<td>Bio Rad</td>
			<td>1845025</td>
			<td>Used for genotyping the mosquito DNA samples and analyzing the thermal denaturation properties of double-stranded DNA (see protocol step 3.3)</td>
		</tr>
		<tr>
			<td>SeqMan Pro</td>
			<td>DNAstar Lasergene software</td>
			<td></td>
			<td>For multiple sequence alignment</td>
		</tr>
		<tr>
			<td>Single-channel pipettor</td>
			<td>Gilson</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tweezers Dumont #5 11 cm</td>
			<td>WPI</td>
			<td>14098</td>
			<td></td>
		</tr>
		<tr>
			<td>White foam plugs</td>
			<td>VWR</td>
			<td>60882-189</td>
			<td></td>
		</tr>
		<tr>
			<td>Wide Drosophila Vials, Polystyrene</td>
			<td>Genesee</td>
			<td>32-117</td>
			<td></td>
		</tr>
		<tr>
			<td>Wide Fly Vial Tray, Blue</td>
			<td>Genesee</td>
			<td>59-164B</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Mosquitoes containing mutations in the genes AaeZIP11 (putative iron transporter21) and myo-fem (a female-biased myosin gene related to flight muscles13) were obtained using CRISPR/Cas9 technology, genotyped using HRMA, and sequence-verified (Figure 5). Figure 5A and Figure 5C show the normalized fluorescence intensity from the HRM curves from AaeZIP11 and <e...

Watch this Scientific Journal Video about Indel Detection following CRISPR/Cas9 Mutagenesis using High-resolution Melt Analysis in the Mosquito Aedes aegypti at JoVE.com

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 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 ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Genetics

3.2K Views.  Texas A&M University. 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. 

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

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Forward genetic screening in model organisms is the workhorse to discover functionally important genes and pathways in many biological processes. In most mutagenesis-based screens, researchers have relied on the use of toxic chemicals, carcinogens, or irradiation, which requires designated equipment, safety setup, and/or disposal of hazardous materials. We have developed a simple approach to induce heritable mutations in C. elegans using germline-expressed histone-miniSOG, a light-inducible potent generator of reactive oxygen species. This mutagenesis method is free of toxic chemicals and requires minimal laboratory safety and waste management. The induced DNA modifications include single-nucleotide changes and small deletions, and complement those caused by classical chemical mutagenesis. This methodology can also be used to induce integration of extrachromosomal transgenes. Here, we provide the details of the LED setup and protocols for standard mutagenesis and transgene integration.

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Genetically-encoded histone-miniSOG induces genome-wide heritable mutations in a blue&#160;light-dependent manner. This mutagenesis method is simple, fast, free of toxic chemicals, and well-suited for forward genetic screening and transgene integration.

Forward genetic screening in model organisms is the workhorse to discover functionally important genes and pathways in many biological processes. In most ...

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 ...

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

In our efforts to determine the patterns of expression and subcellular localization of Drosophila RNAs on a genome-wide basis, and in a variety of tissues, we have developed numerous modifications and improvements to our original fluorescent in situ hybridization (FISH) protocol. To facilitate throughput and cost effectiveness, all steps, from probe generation to signal detection, are performed using exon 96-well microtiter plates. Digoxygenin (DIG)-labelled antisense RNA probes are produced using either cDNA clones or genomic DNA as templates. After tissue fixation and permeabilization, probes are hybridized to transcripts of interest and then detected using a succession of anti-DIG antibody conjugated to biotin, streptavidin conjugated to horseradish peroxidase (HRP) and fluorescently conjugated tyramide, which in the presence of HRP, produces a highly reactive intermediate that binds to electron dense regions of immediately adjacent proteins. These amplification and localization steps produce a robust and highly localized signal that facilitates both cellular and subcellular transcript localization. The protocols provided have been optimized to produce highly specific signals in a variety of tissues and developmental stages. References are also provided for additional variations that allow the simultaneous detection of multiple transcripts, or transcripts and proteins, at the same time.

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

The described RNA in situ hybridization protocol allows the detection of RNA in whole Drosophila embryos or dissected tissues. Using 96-well microtiter plates and tyramide signal amplification, transcripts can be detected at high resolution, sensitivity, and throughput, and at a relatively low cost.

In our efforts to determine the patterns of expression and subcellular localization of Drosophila RNAs on a genome-wide basis, and in a variety of ...

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 ...

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been co-opted as a powerful tool for precise eukaryotic genome engineering. Here, we present a rapid and simple method using chimeric single guide RNAs (sgRNA) and CRISPR-Cas9 Ribonucleoproteins (RNPs) for the efficient and precise generation of genomic point mutations in C. elegans. We describe a pipeline for sgRNA target selection, homology-directed repair (HDR) template design, CRISPR-Cas9-RNP complexing and delivery, and a genotyping strategy that enables the robust and rapid identification of correctly edited animals. Our approach not only permits the facile generation and identification of desired genomic point mutant animals, but also facilitates the detection of other complex indel alleles in approximately 4 - 5 days with high efficiency and a reduced screening workload.

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Here, we present a method to engineer the genome of C. elegans using CRISPR-Cas9 ribonucleoproteins and homology dependent repair templates.

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been ...

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 ...

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Trypanosoma cruzi is a pathogenic protozoan parasite that causes Chagas&#8217; disease mainly in Latin America. In order to identify a novel drug target against T. cruzi, it is important to validate the essentiality of the target gene in the mammalian stage of the parasite, the amastigote. Amastigotes of T. cruzi replicate inside the host cell; thus, it is difficult to conduct a knockout experiment without going through other developmental stages. Recently, our group reported a growth condition in which the amastigote can replicate axenically for up to 10 days without losing its amastigote-like properties. By using this temporal axenic amastigote culture, we successfully introduced gRNAs directly into the Cas9-expressing amastigote to cause gene knockouts and analyzed their phenotypes exclusively in the amastigote stage. In this report, we describe a detailed protocol to produce in vitro derived extracellular amastigotes, and to utilize the axenic culture in a CRISPR/Cas9-mediated knockout experiment. The growth phenotype of knockout amastigotes can be evaluated either by cell counts of the axenic culture, or by replication of intracellular amastigote after host cell invasion. This method bypasses the parasite stage differentiation normally involved in producing a transgenic or a knockout amastigote. Utilization of the temporal axenic amastigote culture has the potential to expand the experimental freedom of stage-specific studies in T. cruzi.

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Here, we describe a protocol to introduce a gene knockout into the extracellular amastigote of Trypanosoma cruzi, using the CRISPR/Cas9 system. The growth phenotype can be followed up either by cell counting of axenic amastigote culture or by proliferation of intracellular amastigotes after host cell invasion.

Trypanosoma cruzi is a pathogenic protozoan parasite that causes Chagas&#8217; disease mainly in Latin America. In order to identify a novel drug target ...

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 ...

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding sites for transcriptional regulators to modulate gene expression. Alterations of CREs have been implicated in various diseases including cancer. While promoters and enhancers have been the primary CREs for studying gene regulation, very little is known about the role of silencer, which is another type of CRE that mediates gene repression. Originally identified as an adaptive immunity system in prokaryotes, CRISPR/Cas9 has been exploited to be a powerful tool for eukaryotic genome editing. Here, we present the use of this technique to delete an intronic silencer in the human RUNX1 gene and investigate the impacts on gene expression in OCI-AML3 leukemic cells. Our approach relies on electroporation-mediated delivery of two preassembled Cas9/guide RNA (gRNA) ribonucleoprotein (RNP) complexes to create two double-strand breaks (DSBs) that flank the silencer. Deletions can be readily screened by fragment analysis. Expression analyses of different mRNAs transcribed from alternative promoters help evaluate promoter-dependent effects. This strategy can be used to study other CREs and is particularly suitable for hematopoietic cells, which are often difficult to transfect with plasmid-based methods. The use of a plasmid- and virus-free strategy allows simple and fast assessments of gene regulatory functions.

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

Direct delivery of preassembled Cas9/guide RNA ribonucleoprotein complexes is a fast and efficient means for genome editing in hematopoietic cells. Here, we utilize this approach to delete a RUNX1 intronic silencer and examine the transcriptional responses in OCI-AML3 leukemic cells.

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding ...

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

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Chapters in this video

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

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

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

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

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

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

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

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

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

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells