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>

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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 will also depend on the time to receive any needed sequencing results (for identification of the SNPs), sequence analyses, and primer ordering. 
The protocol presented here details the step-by-step methods for successful analyses. One of the most critical steps in this process is the proper design of the primers. HRMA is sufficiently sensitive that the presence of even a single SNP within the fragment amplified can result in a strong difference curve that could be misinterpreted as a mutation. Sequencing laboratory colony samples prior to CRISPR mutagenesis allows for the detection and avoidance of SNP-rich regions when designing the target; the HRMA primers will help to prevent this issue. Designing primers for small amplicons (70-100 bp) can also improve the sensitivity and reproducibility of the experiment as larger fragments can present several melt domains, decreasing the chance to distinguish variants22. A limitation of the technique is that completely avoiding SNPs might not be feasible depending on the gene; thus, sequencing and further use of multiple wild-type control individuals may aid the analyses. 
Specifically for this protocol, it is important to keep genotyped mosquitoes alive so they can be backcrossed for the establishment of mutant lines. Removing one of the mosquito legs to extract gDNA may be the least invasive way of achieving that; however, the amount of gDNA recovered from this tissue is minimal. The kit suggested in the Table of Materials was sufficient for obtaining DNA from a mosquito leg and performing the PCR to complete HRMA.
HRMA can offer more flexibility in terms of detection of a broad range of indel sizes, varying from single-base mismatches to multiple base pairs (AaeZIP11 had a 56 bp deletion) and surpassing methods such as the SNA or T7E1 (both enzyme mismatch cleavage assays) that are limited to single-base mismatch or small deletions (~20 bp)5,23. In addition, SNA does not detect homozygous mutants as the PCR products do not contain mismatches. Moreover, SNA results may also be masked by naturally occurring allelic polymorphisms when the nuclease-treated PCR products have a similar length to the mutant alleles. Neither of these limitations is applicable to HRMA. Ultimately, sequencing is necessary to identify the mutant DNA sequence. However, once a mutation in question is known, HRMA used for the identification of mutant individuals is much faster than direct sequencing. This method preserves valuable samples (more mosquitoes will survive, the less they have to wait for their genotype results) and reduces the need for sequencing of large numbers of samples, lowering the total cost of the experiment.

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><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&amp;nbsp; 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 myo-fem mutant samples, respectively, along with wild-type controls. Figure 4B and Figure 4D show the magnification of the difference curves (from the AaeZIP11 and myo-fem mutant samples, respectively) between the melt profiles of the different clusters assigned by the software after subtracting each curve from the wild-type reference. Heterozygous and homozygous mutant AaeZIP11 individuals were placed in different clusters and are easily distinguished from the wild-type controls (Figure 5B). Heterozygous mutant myo-fem individuals are also distinct from the controls (Figure 5D). Note that two clusters within the wild-type controls were present in both cases, most likely due to the presence of SNPs in the target region (Figure 5), highlighting the need to use multiple control samples to avoid categorizing controls as mutants when SNPs cannot be avoided.
Figure 4A shows the sequence analysis of the AaeZIP11 mutant. The electropherogram from AaeZIP11 heterozygous mutants indicates the nucleotide position where the indel occurred. This is represented by a shift from single to double peaks as polymorphic positions will show both nucleotides concomitantly (Figure 4A). The number of base pairs deleted or inserted was calculated by counting the single peaks at the end of the run (Figure 4B) as one of the DNA strands will be shorter or longer than the other by the number of base pairs deleted or inserted, respectively. It is recommended to use sequence-verified gDNA from the mutants as a reference for the identification of the heterozygotes to help with HRMA analyses for subsequent generations. Manual assignment for the curves may be needed when similar curves are automatically assigned to different clusters. This can be done by comparing the temperature-shifted curves and difference curves. Manual adjustment of the software might be needed to properly assign samples to clusters. HRMA results from AaeZIP11 and a mutant called Aeflightin show that individual sample analyses were needed to successfully categorize heterozygotes, homozygotes, and trans-heterozygotes. Initially, the automatic cluster assignment from the software could not make a proper distinction between the groups (Figure 6A, Figure 6C, Figure 6E, and Figure 6G). Each sample was then analyzed individually and assigned to the correct groups based on similarity to reference samples of heterozygotes, homozygotes, and trans-heterozygotes (previously verified by sequencing) (Figure 6B, Figure 6D, Figure 6F, and Figure 6H).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63008/63008fig01.jpg" /> 
Figure 1: SNP identification. Schematic representation of multiple sequence alignment of AaeZIP11 fragment from wild-type. In red are the SNPs, and in green are the fragments free of SNPs; this SNP-free region is suggested for sgRNA and primer design. Abbreviations: SNP = single nucleotide polymorphism; sgRNA = single guide RNA; LVP = Liverpool strain. <a href="https://www.jove.com/files/ftp_upload/63008/63008fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63008/63008fig02.jpg" /> 
Figure 2: Experimental procedure for obtaining genomic DNA from mosquito legs for HRMA. (A) Materials for obtaining genomic DNA including pipettes, tips, PCR plate, optical seal, reservoir, the dilution buffer, and DNA release solution. (B) PCR plate preparation containing DNA release reagent and dilution buffer. (C) Mosquito anesthesia with CO2. (D) Wiping the tweezers with 70% ethanol to prevent contamination between samples. (E) Experimental setup including tweezers, rack for mosquito vials, Petri dish on top of an ice container with anesthetized mosquitoes, and the previously prepared PCR plate. (F) Removal of the mosquito leg. (G) Zoom view of the mosquito leg being submerged in the DNA release reagent solution. (H) Single mosquito placed in the vial. (I) Sealing the PCR plate containing the mosquito legs. Abbreviation: HRMA = high-resolution melt analysis. <a href="https://www.jove.com/files/ftp_upload/63008/63008fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63008/63008fig03.jpg" /> 
Figure 3:&#160;Experimental procedure for HRM analyses. (A) Transferring the released gDNA to a 96-well plate containing the PCR mix. (B) Visual inspection of the difference curves. (C) Marking each sample on the 96-well template with the same color as its respective difference curve color. (D) Backcrossing the individuals from the same cluster. Abbreviations: HRM = high-resolution melt; gDNA = genomic DNA. <a href="https://www.jove.com/files/ftp_upload/63008/63008fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63008/63008fig04.jpg" /> 
Figure 4:&#160;Sequence analysis of an AaeZIP11 mutant. (A) Nucleotide alignment of the AaeZIP11&#916;56 mutant. Dashes highlighted in yellow are the deleted bases. In the box, the electropherogram transitions from single peaks to double peaks, depicting the position where the deletion occurred (arrow). (B) Electropherogram of the end of the sequencing run and the transition from double peaks to single peaks. Note that the number of single peaks represents the number of bases deleted (gray rectangle). Primer sequences are provided in the Supplemental Table S1. <a href="https://www.jove.com/files/ftp_upload/63008/63008fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63008/63008fig05.jpg" /> 
Figure 5:&#160;HRMA of DNA extracted from mosquito legs. DNA was extracted from a single leg from AaeZIP11 (A and B) and myo-fem (C and D) knockout Ae. aegypti mosquitoes and analyzed by HRMA. A and C denote the normalized fluorescence signals of the samples to relative values of 1.0 to 0. B and D denote the magnification of the curve differences by subtracting each curve from the wild-type reference (Liverpool strain). Abbreviations: HRMA = high-resolution melt analysis; LVP = Liverpool strain; RFU = relative fluorescence units. Primer sequences are provided in the Supplemental Table S1. <a href="https://www.jove.com/files/ftp_upload/63008/63008fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/63008/63008fig06.jpg" /> 
Figure 6:&#160;Examples of manual group assignments. (A-D) HRMA for Aeflightin mutants. (A and C) Melt curves (normalized linear scale curves and difference curves, respectively) are automatically grouped by the Precision Melt Analysis Software. (B) Normalization of differential curves altered manually. (D) After altering the normalization of the differential curves, each sample was assigned individually by the peak temperatures in the difference curves for corresponding positive controls (previously sequenced samples identified by &#916;4 and &#916;5 heterozygotes and &#916;4&#916;5 trans-heterozygotes), enabling the clear identification of the 3 groups. Red and brown arrows are added to highlight the peaks at different temperatures. (E-H) HRMA for AaeZIP11 mutants. (E and G) Melt curves are automatically assigned by the software. (F and H) Mutants in the second heterozygous self-cross were appropriately assigned to the correct groups by the similarity of the normalized and difference curves to previously determined references (color-coded in curves). Heterozygotes asg, Homozygotes asg, and Trans-heterozygotes asg: assigned melt curves by Precision Melt Analysis Software. Abbreviations: HRMA = high-resolution melt analysis; LVP = Liverpool strain; RFU = relative fluorescence units. Primer sequences are provided in Supplemental Table S1. <a href="https://www.jove.com/files/ftp_upload/63008/63008fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Material: Detailed protocol for setting up HRMA in the CFX96 Real-Time System (e.g., Bio-rad). <a href="https://www.jove.com/files/ftp_upload/63008/Supplemental material_R2.docx" target="_blank">Please click here to download this File.</a>
Supplemental Figure S1:&#160;Step-by-step instructions for setting up the cycling protocol on Bio-rad CFX Manager. See Supplemental Material 2.1-2.3. <a href="https://www.jove.com/files/ftp_upload/63008/8 - Supplemental Figure 1.png" target="_blank">Please click here to download this File.</a>
Supplemental Figure S2: Step-by-step instructions for a plate setup on Bio-rad CFX Manager. See Supplemental Material 3.1-3.4. <a href="https://www.jove.com/files/ftp_upload/63008/9 - Supplemental Figure 2.png" target="_blank">Please click here to download this File.</a>
Supplemental Figure S3: Step-by-step instructions for a plate setup on Bio-rad CFX Manager. See Supplemental Material 3.5-3.6. <a href="https://www.jove.com/files/ftp_upload/63008/10 - Supplemental Figure 3.png" target="_blank">Please click here to download this File.</a>
Supplemental Figure S4: Step-by-step instructions for the run setup on Bio-rad CFX Manager. See Supplemental Material 4.1. <a href="https://www.jove.com/files/ftp_upload/63008/11 - Supplemental Figure 4.png" target="_blank">Please click here to download this File.</a>
Supplemental Figure S5: Step-by-step instructions for HRMA analysis on Bio-rad CFX Manager. See Supplemental Material 5.1-5.3. Abbreviation: HRMA = high-resolution melt analysis. <a href="https://www.jove.com/files/ftp_upload/63008/12 - Supplemental Figure 5.png" target="_blank">Please click here to download this File.</a>
Supplemental Table S1: List of primers. <a href="https://www.jove.com/files/ftp_upload/63008/Supplemental Table S1.xlsx" target="_blank">Please click here to download this Table.</a>

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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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

Using a Fluorescent PCR-capillary Gel Electrophoresis Technique to Genotype CRISPR/Cas9-mediated Knockout Mutants in a High-throughput Format

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play in the functioning of cells and whole organisms. This cause has been tremendously aided by the recent introduction of the CRISPR/Cas9 system-a versatile tool that allows researchers to manipulate the genome and transcriptome in order to, among other things, knock out, knock down, or knock in genes in a targeted manner. For the purpose of knocking out a gene, CRISPR/Cas9-mediated double-strand breaks recruit the non-homologous end-joining DNA repair pathway to introduce the frameshift-causing insertion or deletion of nucleotides at the break site. However, an individual guide RNA may cause undesirable off-target effects, and to rule these out, the use of multiple guide RNAs is necessary. This multiplicity of targets also means that a high-volume screening of clones is required, which in turn begs the use of an efficient high-throughput technique to genotype the knockout clones. Current genotyping techniques either suffer from inherent limitations or incur high cost, hence rendering them unsuitable for high-throughput purposes. Here, we detail the protocol for using fluorescent PCR, which uses genomic DNA from crude cell lysate as a template, and then resolving the PCR fragments via capillary gel electrophoresis. This technique is accurate enough to differentiate one base-pair difference between fragments and hence is adequate in indicating the presence or absence of a frameshift in the coding sequence of the targeted gene. This precise knowledge effectively precludes the need for a confirmatory sequencing step and allows users to save time and cost in the process. Moreover, this technique has proven to be versatile in genotyping various mammalian cells of various tissue origins targeted by guide RNAs against numerous genes, as shown here and elsewhere.

The genotyping technique described here, which couples fluorescent polymerase chain reaction (PCR) to capillary gel electrophoresis, allows for high-throughput genotyping of nuclease-mediated knockout clones. It circumvents limitations faced by other genotyping techniques and is more cost effective than sequencing methods.

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play ...

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point mutations, which often are responsible for a variety of human inherited disorders. Using a well-established cell-based model system, the point mutation of a single-copy mutant eGFP gene integrated into HCT116 cells has been repaired using this combinatorial approach. The analysis of corrected and uncorrected cells reveals both the precision of gene editing and the development of genetic lesions, when indels are created in uncorrected cells in the DNA sequence surrounding the target site. Here, the specific methodology used to analyze this combinatorial approach to the gene editing of a point mutation, coupled with a detailed experimental strategy to measuring indel formation at the target site, is outlined. This protocol outlines a foundational approach and workflow for investigations aimed at developing CRISPR/Cas9-based gene editing for human therapy. The conclusion of this work is that on-site mutagenesis takes place as a result of CRISPR/Cas9 activity during the process of point mutation repair. This work puts in place a standardized methodology to identify the degree of mutagenesis, which should be an important and critical aspect of any approach destined for clinical implementation.

This protocol outlines the workflow of a CRISPR/Cas9-based gene editing system for the repair of point mutations in mammalian cells. Here, we use a combinatorial approach to gene editing with a detailed follow-on experimental strategy for measuring indel formation at the target site&#8212;in essence, analyzing onsite mutagenesis.

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point ...

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

Genome Editing in Mammalian Cell Lines using CRISPR-Cas

The clustered regularly interspaced short palindromic repeats (CRISPR) system functions naturally in bacterial adaptive immunity, but has been successfully repurposed for genome engineering in many different living organisms. Most commonly, the wildtype CRISPR associated 9 (Cas9) or Cas12a endonuclease is used to cleave specific sites in the genome, after which the DNA double-stranded break is repaired via the non-homologous end joining (NHEJ) pathway or the homology-directed repair (HDR) pathway depending on whether a donor template is absent or present respectively. To date, CRISPR systems from different bacterial species have been shown to be capable of performing genome editing in mammalian cells. However, despite the apparent simplicity of the technology, multiple design parameters need to be considered, which often leave users perplexed about how best to carry out their genome editing experiments. Here, we describe a complete workflow from experimental design to identification of cell clones that carry desired DNA modifications, with the goal of facilitating successful execution of genome editing experiments in mammalian cell lines. We highlight key considerations for users to take note of, including the choice of CRISPR system, the spacer length, and the design of a single-stranded oligodeoxynucleotide (ssODN) donor template. We envision that this workflow will be useful for gene knockout studies, disease modeling efforts, or the generation of reporter cell lines.

CRISPR-Cas is a powerful technology to engineer the complex genomes of plants and animals. Here, we detail a protocol to efficiently edit the human genome using different Cas endonucleases. We highlight important considerations and design parameters to optimize editing efficiency.

The clustered regularly interspaced short palindromic repeats (CRISPR) system functions naturally in bacterial adaptive immunity, but has been ...

Site-Directed &#966;C31-Mediated Integration and Cassette Exchange in Anopheles Vectors of Malaria

Functional genomic analysis and related strategies for genetic control of malaria rely on validated and reproducible methods to accurately modify the genome of Anopheles mosquitoes. Amongst these methods, the &#966;C31 system allows precise and stable site-directed integration of transgenes, or the substitution of integrated transgenic cassettes via recombinase-mediated cassette exchange (RMCE). This method relies on the action of the Streptomyces &#966;C31 bacteriophage integrase to catalyze recombination between two specific attachment sites designated attP (derived from the phage) and attB (derived from the host bacterium). The system uses one or two attP sites that have been integrated previously into the mosquito genome and attB site(s) in the donor template DNA. Here we illustrate how to stably modify the genome of attP-bearing Anopheles docking lines using two plasmids: an attB-tagged donor carrying the integration or exchange template and a helper plasmid encoding the &#966;C31 integrase. We report two representative results of &#966;C31-mediated site-directed modification: the single integration of a transgenic cassette in An. stephensi and RMCE in An. gambiae mosquitoes. &#966;C31-mediated genome manipulation offers the advantage of reproducible transgene expression from validated, fitness neutral genomic sites, allowing comparative qualitative and quantitative analyses of phenotypes. The site-directed nature of the integration also substantially simplifies the validation of the single insertion site and the mating scheme to obtain a stable transgenic line. These and other characteristics make the &#966;C31 system an essential component of the genetic toolkit for the transgenic manipulation of malaria mosquitoes and other insect vectors.

The protocol describes how to achieve site-directed modifications in the genome of Anopheles malaria mosquitoes using the &#966;C31 system. Modifications described include both the integration and the exchange of transgenic cassettes in the genome of attP-bearing docking lines.

Functional genomic analysis and related strategies for genetic control of malaria rely on validated and reproducible methods to accurately modify the ...

Digital-Droplet PCR to Detect Indels Mutations in Genetically Modified Anopheline Mosquito Populations

Recent advances in mosquito genomics and genetic engineering technologies have fostered a need for quick and efficient methods for detecting targeted DNA sequence variation on a large scale. Specifically, detecting insertions and deletions (indels) at gene-edited sites generated by CRISPR guide RNA (gRNA)/Cas9-mediated non-homologous end-joining (NHEJ) is important for assessing the fidelity of the mutagenesis and the frequency of unintended changes. We describe here a protocol for digital-droplet PCR (ddPCR) that is well-suited for high-throughput NHEJ analysis. While this method does not produce data that identifies individual sequence variation, it provides a quantitative estimate of the sequence variation within a population. Additionally, with appropriate resources, this protocol can be implemented in a field-site laboratory setting more easily than next-generation or Sanger sequencing. ddPCR also has a faster turn-around time for results than either of those methods, which allows a more quick and complete analysis of genetic variation in wild populations during field trials of genetically-engineered organisms.

This protocol provides the steps from DNA extraction to experimental set-up for digital droplet PCR (ddPCR), including analysis for the identification and quantification of non-homologous end-joining (NHEJ) events at target sites following gRNA-induced Cas9 cleavage and DNA repair. Other uses of this method include applications such as polymorphism detection and gene-editing variant verification.

Recent advances in mosquito genomics and genetic engineering technologies have fostered a need for quick and efficient methods for detecting targeted DNA ...

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

The emergence of the clustered, regularly interspersed, short palindromic repeats (CRISPR)-Cas9 technology has revolutionized the genetic engineering field and opened the doors for precise genome editing in multiple species, including non-model organisms. In the mosquito Aedes aegypti, loss-of-function mutations and DNA insertions have been accomplished with this technology. Here, we describe a detailed protocol for genome editing through embryonic microinjection in the mosquito A. aegypti using the CRISPR-Cas9 technology, focusing on both the generation of gene knockout and knockin lines. In this protocol, quartz needles are filled with a mixture of guide RNA, recombinant Cas9, and a plasmid containing a DNA cassette encoding a gene for a fluorescent marker, if gene knockin is desired. Embryos at the preblastoderm stage are lined up onto a strip of double-sided sticky tape placed onto a coverslip, which is subsequently mounted onto a glass slide. With the help of a microinjector, the needles are inserted gently into the posterior end of the embryos and a small volume of the CRISPR mixture is dispensed. When the embryos are hatched, the larvae are checked under the fluorescent scope, and the pupae are sex-sorted and separated in different cages. Once the adults emerge, these are reciprocally crossed with wild-type individuals, blood-fed, and placed for egg laying. Once these eggs are hatched, the fluorescent larvae collected represent individuals with stable insertion of the DNA cassette into their genome. These larvae are then grown to the adult stage, outcrossed to wild-type individuals, and then further assessed through molecular techniques to confirm that the exact sequence of the DNA cassette is present at the desired site of the mosquito genome. Homozygous lines can also be obtained by following the provided pipeline of crossing schema and molecular screening of the mutations.

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

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

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

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

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

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

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

Behavior

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

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

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

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

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

Biology

Digital Droplet Polymerase Chain Reaction for Indel Mutation Detection in Target Genes

Source: Carballar-Lejaraz&#250;, R., et al. <a href="https://app.jove.com/v/62607">Digital-Droplet PCR to Detect Indels Mutations in Genetically Modified Anopheline Mosquito Populations.</a> J. Vis. Exp. (2021).
This video demonstrates the role of digital droplet polymerase chain reaction in identifying indel mutation in a DNA sample. The high precision and sensitivity of ddPCR enable accurate and consistent detection of rare mutations present in a sample, even at low levels. It can identify and quantify indel mutations in different regions of the genome, providing valuable insights into the mechanisms of mutation and the functional consequences of these mutations.&#160;

Source: Carballar-Lejaraz&#250;, R., et al. Digital-Droplet PCR to Detect Indels Mutations in Genetically Modified Anopheline Mosquito Populations. J. ...

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

Summary

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Chapters in this video

Using a Fluorescent PCR-capillary Gel Electrophoresis Technique to Genotype CRISPR/Cas9-mediated Knockout Mutants in a High-throughput Format

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Genome Editing in Mammalian Cell Lines using CRISPR-Cas

Site-Directed φC31-Mediated Integration and Cassette Exchange in Anopheles Vectors of Malaria

Digital-Droplet PCR to Detect Indels Mutations in Genetically Modified Anopheline Mosquito Populations

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

Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera (Hübner)

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

Digital Droplet Polymerase Chain Reaction for Indel Mutation Detection in Target Genes