Funding was provided by the University of California Irvine Malaria Initiative. AAJ is a Donald Bren Professor at the University of California, Irvine.

1. DNA extraction
<ol>
	<li>Prepare EDTA/Nuclei Lysis Buffer (EDTA/NLS) with the ratio of 500 &#181;L of NLS and 120 &#181;L of EDTA per sample. Scale-up for multiple samples. Chill the mixture on ice. 
	NOTE: The solution will turn cloudy in 2-5 min when chilled depending on the volume.</li>
	<li>Homogenize the mosquito sample using a mechanical homogenizer for 10-15 s in a 1.5 mL microcentrifuge tube filled with 600 &#181;L of chilled EDTA/NLS; mix thoroughly.</li>
	<li>Add 17.5 &#181;L of 20 mg/mL of Proteinase K to the tube and mix thoroughly.</li>
	<li>Incubate overnight at 55 &#176;C. Alternately, incubate the sample at 55 &#176;C for 3 h with shaking and vortex the sample every 1 h.</li>
	<li>Add 200 &#181;L of Protein Precipitation Solution to the room temperature sample and vortex vigorously for 20 s.</li>
	<li>Chill the sample for 5 min on ice.</li>
	<li>Centrifuge the sample to pellet proteins at 15,890 RCF for 4 min.</li>
	<li>Carefully aspirate the supernatant which contains the DNA and transfer it to a clean 1.5 mL microcentrifuge tube containing 600 &#181;L of isopropanol.</li>
	<li>Gently mix the solution by inverting the tube 5-10 times. Centrifuge for 1 min at 15,890 RCF. Carefully decant the supernatant while preserving the DNA pellet.</li>
	<li>Add 600 &#181;L of room-temperature 70% ethanol. Wash the pelleted DNA by gently inverting the tube.</li>
	<li>Centrifuge for 1 min at 15,890 RCF. Carefully remove the supernatant by aspiration using a glass pipette tip.</li>
	<li>Invert the tube onto clean absorbent paper and air-dry the pellet for 10-15 min.</li>
	<li>Resuspend the DNA with PCR-grade water. Use 20 &#181;L per individual mosquito sample or 100 &#181;L for 10 pooled mosquitoes. 
	&#8203;NOTE: DNA extraction methods for mosquito samples using a commercially-available kit (see Table of Materials) are adapted from the manufacturer&#39;s Isolating Genomic DNA from Tissue Culture Cells and Animal Tissue protocol.</li>
</ol>
2. ddPCR reactions and droplet generation preparation
<ol>
	<li>Quantify DNA using a fluorometer. 
	NOTE: For the drop-off assay, it is recommended to use a range of 3,000-30,000 haploid genome copies per reaction, which is designed to detect NHEJ events with a HEX-labeling probe that binds to a WT sequence of the targeted cut site and will not anneal (drop-off) if there is a deletion or insertion at the target site, indicating the presence of an NHEJ variant.</li>
	<li>Calculate the copy number using the haploid genome weight and concentration of DNA in the extract. This is done by multiplying the concentration of the extracted DNA by the volume used to obtain the total DNA mass, then dividing it by the haploid genome weight. Ensure that the volume added is between 1-10 &#181;L. Dilute as necessary to be in the recommended haploid genome copy range. 
	NOTE: One Anopheles gambiae haploid genome is estimated to be 0.27 pg per adult mosquito10.</li>
	<li>Design primers and probes. Design forward and reverse oligonucleotide primers with a Primer Melting Temperature (Tm) in the range of 55-60 &#176;C that flank the 5&#39;- and 3&#39;-ends of the gRNA target site producing an amplicon of 150-400 bp.
	<ol>
		<li>HEX (Hexachloro-fluorescein)-labeled probe for NHEJ detection: Design an oligonucleotide of ~15-20 bp in length complementary to the target site and add the HEX-probe to the 5&#39;-end and BHQ1 (BLack Hole Quencher 1) to the 3&#39; end. The Tm of the hydrolysis probe should be 3-10&#160;&#176;C higher than the Tm of the primers.</li>
		<li>FAM (6-carboxyfluorescein)-labeled probe for reference WT: Design an oligonucleotide ~15-20 bp in length complementary to a conserved genome site distant (about 25 bp) from the target site and add the FAM-probe to the 5&#39;-end and BHQ1 to the 3&#39; end. The Tm of the hydrolysis probe should be 3-10&#160;&#176;C higher than the Tm of the primers.</li>
	</ol>
	</li>
</ol>
3. PCR reaction preparation
<ol>
	<li>Prepare 25 &#181;L of the ddPCR Sample Mix with the following components: ddPCR supermix for probes (no UTP): 12.5 &#181;L, forward and reverse primers (10 &#181;M): 1 &#181;L each, HEX/FAM probes (10 &#181;M): 0.625 &#181;L each, DNA: 1-5 &#181;L (3,750-37,500 haploid genome copies) and water: up to 25 &#181;L.</li>
	<li>Thoroughly mix the reactions by vortexing or reflux pipetting (up and down) (20x). 
	NOTE: If the reactions are in a 96-well plate, pipette the entire volume up and down 20 times rather than vortexing to avoid bubble formation.</li>
	<li>Briefly centrifuge the samples to settle the mixture at the bottom of the tube or well. 
	&#8203;NOTE: Ensure the reactions are at room temperature for the droplet generation. Prepare 1x of ddPCR mix for extra/unused wells in each cartridge (each cartridge has 8 wells).</li>
</ol>
4. Droplet generation
<ol>
	<li>Using a 50 &#181;L multichannel pipette, load 20 &#181;L of the ddPCR sample mix into the middle row of the cartridge (Figure 1A, top).</li>
	<li>Load 70 &#181;L of the oil into the bottom row. Load 20 &#181;L of 1x ddPCR supermix into unused wells. 
	NOTE: Do not introduce bubbles.</li>
	<li>Place the gasket touching only the edges, avoiding the center concaved area (Figure 1B).</li>
	<li>Place the plate securely in the droplet generator and close the cover to start the run.</li>
	<li>Using the multichannel pipette, transfer 40 &#181;L of the emersion mix from the top row of the cartridge (Figure 1A, bottom) into the 96-well plate.
	<ol>
		<li>Draw liquid sample for 3-5 s at a 45-30&#176; angle. Expel the mixture slowly for over 3 s at a 45&#176; angle into the side of the well, allowing it to drip down the side. It is okay to go to the second stop (complete expulsion) of the pipette to expel all the liquid.</li>
	</ol>
	</li>
	<li>Using foil heat seals, seal the plates for 5 s at 180 &#176;C.</li>
</ol>
5. PCR
<ol>
	<li>Place the sealed plate into the thermocycler (Figure 1C) and set the recommended PCR conditions if following the NHEJ Drop-Off guidelines as follows:
	<ol>
		<li>Initial denaturation at 95 &#176;C for 10 min.</li>
		<li>Set 40 cycles of 94 &#176;C for 30 s to denature, 55 &#176;C for 1 min to anneal, and 60 &#176;C for 2 min to extend.</li>
		<li>Hold at 98 &#176;C for 10 min.</li>
		<li>Hold at 4 &#176;C. 
		&#8203;NOTE: The annealing temperature for specific primers and probe sets may be optimized using a thermal gradient. Use a ramp rate of 2 &#176;C/s for all the steps. PCR conditions should be adjusted depending on each experimental design and set-up.</li>
	</ol>
	</li>
</ol>
6. Droplet reading
<ol>
	<li>Place the plate securely in the droplet reader with well A-1 at the top left (smoothed corner, the other three are edged) (Figure 1D).</li>
	<li>Set up the plate in the program: Designating FAM as the known reference channel and HEX as the unknown one (Figure 2A).</li>
	<li>Run the Droplet reading experiment as direct quantification. After the run finishes, change the Experiment type to Drop-Off (DOF) for the analysis (Figure 2A).</li>
</ol>
7. Analysis
<ol>
	<li>Designate the correct experimental parameters (Figure 2A): Sample Information, SuperMix, Target Name (WT or NHEJ), Target Type (Ref or Unknown), Signal Ch1 (HEX or FAM), Signal Ch2 (HEX or FAM), and manually set the threshold for droplet count (recommend above 10,000 for reliable results). The software will perform most of the analysis with the designated parameter.</li>
	<li>Check the droplet count in the Droplet tab; ensure that all are above 10,000 (Figure 2B).</li>
	<li>Check 1 D amplitude for efficient signal separation from negatives.</li>
	<li>In the Plate Editor, highlight the entire plate and set the Experiment Type to Drop off.</li>
	<li>Set the WT target as a Reference, designating channel one for FAM and channel two for HEX.</li>
	<li>Set NHEJ target as unknown, designating channel one for FAM and channel two as none.</li>
	<li>In the 2D amplitude tab, set the Cluster Thresholds with the graph tools for each sample. Consider the tail associated with the WT cluster; this is normal for NHEJ assays (Figure 3A). 
	NOTE: Under the Ratio tab, click on the Gear Icon from the top-right of the graph. Select the Fractional Abundance. The graph will now plot a point corresponding to the percentage of NHEJ events (Figure 3B).</li>
</ol>

<ol><li>Gantz, V. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Highly+efficient+Cas9-mediated+gene+drive+for+population+modification+of+the+malaria+vector+mosquito+Anopheles+stephensi%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Highly efficient Cas9-mediated gene drive for population modification of the malaria vector mosquito Anopheles stephensi.</a> Proceedings of the National Academy of Sciences of the United States of America. 112 (49), 6736-6743 (2015).</li>
<li>Hammond, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+CRISPR-Cas9+gene+drive+system+targeting+female+reproduction+in+the+malaria+mosquito+vector+Anopheles+gambiae%5BTitle%5D)+AND+%22Nature+Biotechnology%22%5BJournal%5D)">A CRISPR-Cas9 gene drive system targeting female reproduction in the malaria mosquito vector Anopheles gambiae.</a> Nature Biotechnology. 34, 78-83 (2016).</li>
<li>Courtier-Orgogozo, V., Morizot, B., Boëte, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Agricultural+pest+control+with+CRISPR-based+gene+drive%3A+time+for+public+debate%3A+Should+we+use+gene+drive+for+pest+control%5BTitle%5D)+AND+%22EMBO+Reports%22%5BJournal%5D)">Agricultural pest control with CRISPR-based gene drive: time for public debate: Should we use gene drive for pest control.</a> EMBO Reports. 18 (6), 878-880 (2017).</li>
<li>Carballar-Lejarazú, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Next-generation+gene+drive+for+population+modification+of+the+malaria+vector+mosquito%2C+Anopheles+gambiae%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Next-generation gene drive for population modification of the malaria vector mosquito, Anopheles gambiae.</a> Proceedings of the National Academy of Sciences of the United States of America. 117 (37), 22805-22814 (2020).</li>
<li>Adolfi, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Efficient+population+modification+gene-drive+rescue+system+in+the+malaria+mosquito+Anopheles+stephensi%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Efficient population modification gene-drive rescue system in the malaria mosquito Anopheles stephensi.</a> Nature Communications. 11, 5553(2020).</li>
<li>Wang, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Comparison+of+droplet+digital+PCR+and+direct+Sanger+sequencing+for+the+detection+of+the+BRAFV600E+mutation+in+papillary+thyroid+carcinoma%5BTitle%5D)+AND+%22Journal+of+Clinical+Laboratory+Analysis%22%5BJournal%5D)">Comparison of droplet digital PCR and direct Sanger sequencing for the detection of the BRAFV600E mutation in papillary thyroid carcinoma.</a> Journal of Clinical Laboratory Analysis. 33 (6), 22902(2019).</li>
<li>Dong, L., Wang, S., Fu, B., Wang, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Evaluation+of+droplet+digital+PCR+and+next+generation+sequencing+for+characterizing+DNA+reference+material+for+KRAS+mutation+detection%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">Evaluation of droplet digital PCR and next generation sequencing for characterizing DNA reference material for KRAS mutation detection.</a> Scientific Reports. 8 (1), 9650(2018).</li>
<li>Bell, C. C., Magor, G. W., Gillinder, K. R., Perkins, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+high-throughput+screening+strategy+for+detecting+CRISPR_Cas9+induced+mutations+using+next-generation+sequencing%5BTitle%5D)+AND+%22BMC+Genomics%22%5BJournal%5D)">A high-throughput screening strategy for detecting CRISPR_Cas9 induced mutations using next-generation sequencing.</a> BMC Genomics. 15 (1), 1002(2014).</li>
<li>Carballar-Lejarazú, R., Kelsey, A., Pham, T. B., Bennett, E. P., James, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Digital+droplet+PCR+and+IDAA+for+the+detection+of+CRISPR+indel+edits+in+the+malaria+species+Anopheles+stephensi%5BTitle%5D)+AND+%22Biotechniques%22%5BJournal%5D)">Digital droplet PCR and IDAA for the detection of CRISPR indel edits in the malaria species Anopheles stephensi.</a> Biotechniques. 68 (4), 172-179 (2020).</li>
<li>Holt, R. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+genome+sequence+of+the+malaria+mosquito+Anopheles+gambiae%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">The genome sequence of the malaria mosquito Anopheles gambiae.</a> Science. 298 (5591), 129-149 (2002).</li>
</ol>

The authors have no disclosures.

Digital-droplet PCR is an efficient method to determine the presence of indel alleles resulting from NHEJ events in a Cas/gRNA-based gene-drive system and allows for quantification of the frequency of these alleles in individuals or populations. Some steps of the protocol need to be followed with special care to achieve reliable results. Firstly, the genomic DNA extraction needs to be performed carefully to ensure high quality and sufficient quantity. A good extraction will allow accurate determination of the haploid genome copies per reaction. In our experience, a commercially-available kit (see Table of Materials) provided consistently high-quality DNA extractions. However, extractions of individual mosquitoes can prove to be particularly challenging as the DNA pellet becomes hard to visualize and can easily be sucked up with the supernatant if not careful. Secondly, primers and probes must be designed carefully. Before completing the ddPCR experiment, ensure that the designed primers result in a single PCR product by first performing a traditional PCR and visualizing a single product via gel electrophoresis. The reference FAM probe also must be designed so that it is complementary to a highly-conserved sequence. This will ensure accurate detections of WT alleles throughout a diverse population. The primer/probe combinations for each unique experiment will have different thermocycler conditions, and it is recommended to optimize those conditions using a thermal gradient.
Other methods for identifying indels exist, such as Sanger sequencing or NGS. Sanger sequencing is limited because it has a lower limit of detection and low discovery power to identify novel variants. Sanger sequencing also is labor-intensive and is not high-throughput. Compared to Sanger sequencing, NGS does not have the same limitations of low sensitivity, discovery power, and throughput. Another benefit of NGS is its ability to detect a variety of mutations from single nucleotide polymorphism (SNPs) to rearrangements. However, NGS is a more costly and time-consuming method in the application of determining Cas9/gRNA-associated indels because there is only one target region of interest, and it is best suited for larger genome-wide analyses. Compared to the aforementioned methods, ddPCR is high throughput and has a quick turn-around time. If the ddPCR materials and instruments are available in-house, 96 samples can be processed within 1-2 days, making it well suited for quick analysis of large trials of Cas9/gRNA -modified organisms.
While many benefits exist for ddPCR there are also limitations. Firstly, the ddPCR equipment is not frequently available in an independent laboratory environment. ddPCR equipment may be available communally at larger research institutions, but this does not allow for ease of data generation and analysis outside of the institution. Secondly, unlike the alternatives, ddPCR does not provide the individual unique sequences of identified indel mutations. Digital-droplet PCR will provide the frequency of indel mutations within a population, but without the sequence, one cannot determine whether the indels present are more likely to conserve or inhibit the function of the gene of interest. The ddPCR method is perhaps suited best to analyze wild populations after a field release trial of a Cas9/gRNA-based drive organism because it can efficiently determine the frequency of introduction of the transgene into the native population and the generation of indels within the population in close to real-time. Due to the ddPCR quick turn-around time, it would be feasible to perform sampling and analyze the population in a field trial region weekly if materials were available locally. The start-up costs to purchase, import, and set up the ddPCR equipment would be high in remote laboratories but the benefits of being able to assess rigorously a wild population as it is undergoing modification from a drive system would justify the costs.

Gene drives have immense potential to control insect populations of medical and agricultural relevance1,2,3,4,5. For example, gene-drive systems based on CRISPR Cas nucleases and guide RNAs (gRNAs) can be used to modify vector mosquito populations by introducing traits that confer refractoriness to malaria parasites leading to reduced transmission and less disease1,4,5. The gene-drive system copies itself and the associated trait from one homologous chromosome to another in the pre-meiotic germ cells, and this ensures that the majority of the offspring inherit the drive and create the potential for long-lasting and sustainable population modification in the field. However, one disadvantage of the Cas/gRNA-based methods is the possibility of generating insertion and deletion (indel) mutations through non-homologous end-joining (NHEJ) DNA repair, resulting in the generation of drive-resistant alleles, which when accumulated to a high enough frequency in the population, can stop the drive system from spreading1,2,3,4. This protocol details a high-throughput and reliable method that can determine the prevalence and relative quantity of indel mutations, at both the population and individual level, during Cas/gRNA-based gene drive.
Next-generation sequencing (NGS) methods provide unparalleled sequencing resolution. However, the cost and technical requirements associated with NGS prohibit routine testing and limit its use as a high-throughput method to assess indels6,7,8. Traditional PCR quantification methods have long been used as the standard evaluation procedure for genome indels; however, these methods are labor-intensive, take a long time to procure data, and have a high degree of variability. Digital-Droplet PCR (ddPCR) has been proven to be more sensitive at detecting mutations than Sanger sequencing in some applications and has a lower detection limit than NGS in others6,7,8,9. Moreover, the cost to assess a sample set and turn-around time for obtaining results is less expensive and faster, respectively, for ddPCR than either Sanger sequencing or NGS9. Using a dual-probe system, the Drop-Off assay identifies NHEJ alleles based on the absence of wild-type (WT) sequence at the gRNA-directed target Cas9 cut site. In this assay, a short amplicon including the predicted cut site of the Cas/gRNA-based system&#160;is amplified with a specific primer pair. One fluorescent probe is designed to bind to a conserved region of the amplicon and another fluorescent probe recognizes the WT sequence of the cut site. In the presence of an NHEJ allele, the latter will not bind to the amplicon.
The use of ddPCR provides the ability to design primers to target deletions, single base-pair differences and insertions, which will allow for NHEJ profiling in mosquito population analyses9. Given these attractive features, we created a protocol for ddPCR for high-throughput detection of indels generated from a Cas/gRNA-based gene-drive system in mosquitoes.

<table><tbody><tr><td>&lt;strong&gt;Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>ddPCR Super Mix for Probes (no dUTP)</td><td>Bio-Rad</td><td>1863024</td><td></td></tr><tr><td>DNA extraction reagent (e.g. Wizard Genomic DNA Purification kit)</td><td>Promega</td><td>A1120</td><td></td></tr><tr><td>EDTA (pH 8.0)</td><td>Invitrogen</td><td>AM9260G</td><td></td></tr><tr><td>Ethanol, 200 Proof</td><td>Thermo Fisher Scientific</td><td>A4094</td><td></td></tr><tr><td>Isopropanol (Certified ACS)</td><td>Thermo Fisher Scientific</td><td>A416-500</td><td></td></tr><tr><td>Nuclei Lysis Solution (NLS) (Wizard Genomic DNA Purification kit)</td><td>Promega</td><td>A1120</td><td></td></tr><tr><td>PCR-grade Water</td><td></td><td></td><td>Any certified PCR-grade water can be used</td></tr><tr><td>Protein Precipitation Solution (Wizard Genomic DNA Purification kit)</td><td>Promega</td><td>A1120</td><td></td></tr><tr><td>Proteinase K 20 mg/mL</td><td>Thermo Fisher Scientific</td><td>AM2546</td><td></td></tr><tr><td>&lt;strong&gt;Materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>ddPCR 96-Well Plate</td><td>Bio-Rad</td><td>12001925</td><td></td></tr><tr><td>Droplet Generator DG8 Cartridge and Gaskets</td><td>Bio-Rad</td><td>1864007</td><td></td></tr><tr><td>Droplet Generation Oil for probes</td><td>Bio-Rad</td><td>1863005</td><td></td></tr><tr><td>Fluorescent probes (e.g. FAM/HEX probes)</td><td>Sigma-Aldrich</td><td>N/A</td><td>Probes are experiment specific and can be purchased from any certified seller available.</td></tr><tr><td>Forward and Reverse oligonucleotide primers</td><td>Sigma-Aldrich</td><td>N/A</td><td>Primers are experiment specific and can be purchased from any certified seller available.</td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>C1000 Touch Thermal Cycler</td><td>Bio-Rad</td><td>1851148</td><td>Can use other Thermo cycler with gradient function and deep well</td></tr></tbody></table>

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.

An application of this procedure appears in Carballar-Lejaraz&#250; et al.9. The ddPCR Drop-off assay utilizes two fluorescent probes to discern WT and indel sequences: A FAM probe binds to a conserved sequence within the amplicon, whereas the HEX probe targets the WT sequence of the targeted site (Figure 4A). In the presence of an indel, the HEX probe will not bind. Representative results can be found in Figure 2, Table 1, and Table 2 of Carballar-Lejaraz&#250; et al.9. Using this protocol, ddPCR has been proven to detect a wide variety of CRISPR-Cas9 induced NHEJ events and quantify the NHEJ frequency in an individual or pooled sample. Fifteen different pooled samples of 10 mosquitoes each contained various NHEJ alleles (Table 2 of Carballar-Lejaraz&#250; et al.9). These were analyzed with ddPCR using the protocol and parameters presented here. Results from Table 19 show that all 15 samples carried 100% indel alleles as identified by the Drop-off assay (Figure 4B). In another experiment, 11 pooled samples of WT mosquitoes and NHEJ mosquitoes with different NHEJ percentages (0%, 10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, and 100%) were examined with this ddPCR protocol, and the results (Figure 2; Carballar- Lejaraz&#250; et al.9) showed that the identified percentage is close to the compared technique of Indel Detection by Amplicon Analysis (Figure 4C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62607/62607fig01.jpg" /> 
Figure 1: Experimental set-up and procedure. (A) Cartridge preparation for Droplet generation. (Top) Samples are filled in the middle row of the cartridge, while oil is filled in the bottom row. (Bottom) Top row filled with emulsified droplets after droplet generation. (B) Droplet generator with a cartridge filled with sample and covered by a gasket in place. (C) 96-well plate covered with foil seal in a Thermo-Cycler. (D) Droplet Reader with 96-well-plate in place with a metal cover latched over the plate to secure it. <a href="https://www.jove.com/files/ftp_upload/62607/62607fig01large.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/62607/62607fig02.jpg" /> 
Figure 2: Droplet reading. (A) Software interface for droplet reading. Orange boxes show wells with samples. Gray boxes are empty wells. Experimental parameters are set up in the Edit Tools panel (right-hand side). Each sample can be edited by clicking on the respective sample box. Select Drop Off (DOF) for Experimental Type. In sample information, fill in the appropriate information for the sample&#39;s Name and Type, as well as SuperMix. Choose the Basic Drop-Off for the Assay Information. For the WT sample, choose WT for the Target Name, Ref for Target Type, and both FAM and HEX for Signal Ch1 and Ch2, respectively. For NHEJ samples, fill in the appropriate name for the Target Name, choose Unknown for the Target Type, and choose FAM for Signal Ch1. Leave Signal Ch2 at None. (B) Droplet count results for multiple samples. <a href="https://www.jove.com/files/ftp_upload/62607/62607fig02large.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/62607/62607fig03.jpg" /> 
Figure 3:&#160;Drop-Off assay analysis. (A) Cluster 2D plot for the droplet count of the WT and NHEJ alleles. In the 2D Amplitude tab, all droplets are unclassified by default. In this figure, colors are manually assigned for distinguishing. The orange dots cluster are WT allele counts obtained by binding of both FAM and HEX probes at the reference sequence and target site sequence, respectively. Blue dots represent scores of droplets that have FAM binding to the reference sequence but no HEX binding at the target site sequence (hence drop-off of HEX). Gray dots are empty droplets that don&#39;t have either FAM or HEX binding. (B) Ratio/Abundance graphs of NHEJ events. Under the Ratio tab, select Fractional Abundance for a graph with the correspondent percentage of NHEJ events. <a href="https://www.jove.com/files/ftp_upload/62607/62607fig03large.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/62607/62607fig04.jpg" /> 
Figure 4: Application of Drop-Off assay with ddPCR for non-homologous end-joining identification and quantification in the transgenic Anopheles stephensi line, AsMCRkh1. (A) Schematic presentation of the ddPCR Drop-Off assay to detect mutations at a targeted DNA site with a dual-probe system. An amplicon of 150-400 bp is amplified with the forward and reverse primers. A FAM-labeled probe is designed to bind to a conserved sequence of the amplicon, whereas a HEX-labeled probe is designed to bind to the WT gRNA targeted site. (B) Detection of various types of indels with ddPCR. Fifteen pools of 10 AsMCRkh1 mosquitoes each containing various types of indel, including insertion, deletion, and substitution, were analyzed with the ddPCR Drop-Off assay. Details of mutations and sequences can be found in Table 2 and Table S3 of Carballar et al.9. (C) Quantification of NHEJ in mixed samples of AsMCRkh1 and WT mosquitoes with various ratios (10:0, 9:1, 8:2, 7:3, 6:4, 5:5, 4:6, 3:7, 2:8, 1:9, and 0:10) using ddPCR and a compared technique of Indel Detection by Amplicon Analysis9. Images adapted from Carballar-Lejaraz&#250; et al. Biotechniques. 68(4):172-179 (2020)9. <a href="https://www.jove.com/files/ftp_upload/62607/62607fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Primer/Probe</td>
			<td>Sequence (5&#8217; &#187; 3&#8217;)</td>
		</tr>
		<tr>
			<td>ddPCR Forward Primer</td>
			<td>ATGATCAAATGTCGACCG</td>
		</tr>
		<tr>
			<td>ddPCR Reverse Primer</td>
			<td>ACCGTACTGGTTGAACA</td>
		</tr>
		<tr>
			<td>ddPCR HEX Probe (BHQ1)</td>
			<td>[HEX]-TTCTACGGGCAGGGC-[BHQ1]</td>
		</tr>
		<tr>
			<td>ddPCR FAM Probe (BHQ1)</td>
			<td>[6FAM]-CCACGTGGGATCGAAGG-[BHQ1]</td>
		</tr>
		<tr>
			<td colspan="2">HEX: Hexachloro-fluorescein, FAM: 6-carboxyfluorescein, BHQ: Black Hole Quencher</td>
		</tr>
	</tbody>
</table>
Table 1: Sequences of primers and probes.

Watch this Scientific Journal Video about Digital-Droplet PCR to Detect Indels Mutations in Genetically Modified Anopheline Mosquito Populations at JoVE.com

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

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 ...

digital-droplet-pcr-to-detect-indels-mutations-genetically-modified

Research

JoVE Journal

Genetics

3.4K Views.  University of California, Irvine. 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.

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

Transfection and Mutagenesis of Target Genes in Mosquito Cells by Locked Nucleic Acid-modified Oligonucleotides

Plasmodium parasites, the causative agent of malaria, are transmitted through the bites of infected Anopheles mosquitoes resulting in over 250 million new infections each year. Despite decades of research, there is still no vaccine against malaria, highlighting the need for novel control strategies. One innovative approach is the use of genetically modified mosquitoes to effectively control malaria parasite transmission. Deliberate alterations of cell signaling pathways in the mosquito, via targeted mutagenesis, have been found to regulate parasite development 1. From these studies, we can begin to identify potential gene targets for transformation. Targeted mutagenesis has traditionally relied upon the homologous recombination between a target gene and a large DNA molecule. However, the construction and use of such complex DNA molecules for generation of stably transformed cell lines is costly, time consuming and often inefficient. Therefore, a strategy using locked nucleic acid-modified oligonucleotides (LNA-ONs) provides a useful alternative for introducing artificial single nucleotide substitutions into episomal and chromosomal DNA gene targets (reviewed in 2). LNA-ON-mediated targeted mutagenesis has been used to introduce point mutations into genes of interest in cultured cells of both yeast and mice 3,4. We show here that LNA-ONs can be used to introduce a single nucleotide change in a transfected episomal target that results in a switch from blue fluorescent protein (BFP) expression to green fluorescent protein (GFP) expression in both Anopheles gambiae and Anopheles stephensi cells. This conversion demonstrates for the first time that effective mutagenesis of target genes in mosquito cells can be mediated by LNA-ONs and suggests that this technique may be applicable to mutagenesis of chromosomal targets in vitro and in vivo.

Oligonucleotides can be used to site specifically substitute a single nucleotide of transfected target genes in both Anopheles gambiae and Anopheles stephensi cells.

Plasmodium parasites, the causative agent of malaria, are transmitted through the bites of infected Anopheles mosquitoes resulting in over 250 million new ...

Immunology and Infection

A Simple Chelex Protocol for DNA Extraction from Anopheles spp.

Endemic countries are increasingly adopting molecular tools for efficient typing, identification and surveillance against malaria parasites and vector mosquitoes, as an integral part of their control programs1,2,3,4,5. For sustainable establishment of these accurate approaches in operations research to strengthen malaria control and elimination efforts, simple and affordable methods, with parsimonious reagent and equipment requirements are essential6,7,8. Here we present a simple Chelex-based technique for extracting malaria parasite and vector DNA from field collected mosquito specimens.
We morphologically identified 72 Anopheles gambiae sl. from 156 mosquitoes captured by pyrethrum spray catches in sleeping rooms of households within a 2,000 km2 vicinity of the Malaria Institute at Macha. After dissection to separate the head and thorax from the abdomen for all 72 Anopheles gambiae sl. mosquitoes, the two sections were individually placed in 1.5 ml microcentrifuge tubes and submerged in 20 &mu;l of deionized water. Using a sterile pipette tip, each mosquito section was separately homogenized to a uniform suspension in the deionized water. Of the ensuing homogenate from each mosquito section, 10 &mu;l was retained while the other 10 &mu;l was transferred to a separate autoclaved 1.5 ml tube. The separate aliquots were subjected to DNA extraction by either the simplified Chelex or the standard salting out extraction protocol9,10. The salting out protocol is so-called and widely used because it employs high salt concentrations in lieu of hazardous organic solvents (such as phenol and chloroform) for the protein precipitation step during DNA extraction9.
Extracts were used as templates for PCR amplification using primers targeting arthropod mitochondrial nicotinamide adenine dinucleotide dehydrogenase (NADH) subunit 4 gene (ND4) to check DNA quality11, a PCR for identification of Anopheles gambiae sibling species10 and a nested PCR for typing of Plasmodium falciparum infection12. Comparison using DNA quality (ND4) PCR showed 93% sensitivity and 82% specificity for the Chelex approach relative to the established salting out protocol. Corresponding values of sensitivity and specificity were 100% and 78%, respectively, using sibling species identification PCR and 92% and 80%, respectively for P. falciparum detection PCR. There were no significant differences in proportion of samples giving amplicon signal with the Chelex or the regular salting out protocol across all three PCR applications. The Chelex approach required three simple reagents and 37 min to complete, while the salting out protocol entailed 10 different reagents and 2 hr and 47 min' processing time, including an overnight step. Our results show that the Chelex method is comparable to the existing salting out extraction and can be substituted as a simple and sustainable approach in resource-limited settings where a constant reagent supply chain is often difficult to maintain.

A Simple Chelex Protocol for DNA Extraction from Anopheles spp.

A rapid and affordable way to extract quality malaria parasite and vector DNA from mosquito specimens is described. Capitalizing on chelating properties of Chelex resin, the simple method enables genotyping of malaria parasites in mosquito mid-gut and salivary gland phases, as well as molecular identification of the Anopheles sibling species by PCR.

Endemic countries are increasingly adopting molecular tools for  efficient typing, identification and surveillance against malaria parasites and  vector ...

Multiplexed Isothermal Amplification Based Diagnostic Platform to Detect Zika, Chikungunya, and Dengue 1

Zika, dengue, and chikungunya viruses are transmitted by mosquitoes, causing diseases with similar patient symptoms. However, they have different downstream patient-to-patient transmission potentials, and require very different patient treatments. Thus, recent Zika outbreaks make it urgent to develop tools that rapidly discriminate these viruses in patients and trapped mosquitoes, to select the correct patient treatment, and to understand and manage their epidemiology in real time.
Unfortunately, current diagnostic tests, including those receiving 2016 emergency use authorizations and fast-track status, detect viral RNA by reverse transcription polymerase chain reaction (RT-PCR), which requires instrumentation, trained users, and considerable sample preparation. Thus, they must be sent to &#34;approved&#34; reference laboratories, requiring time. Indeed, in August 2016, the Center for Disease Control (CDC) was asking pregnant women who had been bitten by a mosquito and developed a Zika-indicating rash to wait an unacceptable 2 to 4 weeks before learning whether they were infected. We very much need tests that can be done on site, with few resources, and by trained but not necessarily licensed personnel.
This video demonstrates an assay that meets these specifications, working with urine or serum (for patients) or crushed mosquito carcasses (for environmental surveillance), all without much sample preparation. Mosquito carcasses are captured on paper carrying quaternary ammonium groups (Q-paper) followed by ammonia treatment to manage biohazards. These are then directly, without RNA isolation, put into assay tubes containing freeze-dried reagents that need no chain of refrigeration. A modified form of reverse transcription loop-mediated isothermal amplification with target-specific fluorescently tagged displaceable probes produces readout, in 30 min, as a three-color fluorescence signal. This is visualized with a handheld, battery-powered device with an orange filter. Forward contamination is prevented with sealed tubes, and the use of thermolabile uracil DNA glycosylase (UDG) in the presence of dUTP in the amplification mixture.

Current multiplexed diagnostics to detect Zika, chikungunya, and dengue viruses require complex sample preparation and expensive instrumentation, and are difficult to use in low resource environments. We show a diagnostic that uses isothermal amplification with target-specific strand displaceable probes to detect and differentiate these viruses with high sensitivity and specificity.

Zika, dengue, and chikungunya viruses are transmitted by mosquitoes, causing diseases with similar patient symptoms. However, they have different ...

Biochemistry

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. One important application of this method is the development of high-quality physical maps useful for improving the genome assemblies for various organisms. The natural banding pattern of polytene and mitotic chromosomes provides guidance for the precise ordering and orientation of the genomic supercontigs. Among the three mosquito genera, namely Anopheles, Aedes, and Culex, a well-established chromosome-based mapping technique has been developed only for Anopheles, whose members possess readable polytene chromosomes 1. As a result of genome mapping efforts, 88% of the An. gambiae genome has been placed to precise chromosome positions 2,3 . Two other mosquito genera, Aedes and Culex, have poorly polytenized chromosomes because of significant overrepresentation of transposable elements in their genomes 4, 5, 6. Only 31 and 9% of the genomic supercontings have been assigned without order or orientation to chromosomes of Ae. aegypti 7 and Cx. quinquefasciatus 8, respectively. Mitotic chromosome preparation for these two species had previously been limited to brain ganglia and cell lines. However, chromosome slides prepared from the brain ganglia of mosquitoes usually contain low numbers of metaphase plates 9. Also, although a FISH technique has been developed for mitotic chromosomes from a cell line of Ae. aegypti 10, the accumulation of multiple chromosomal rearrangements in cell line chromosomes 11 makes them useless for genome mapping. Here we describe a simple, robust technique for obtaining high-quality mitotic chromosome preparations from imaginal discs (IDs) of 4th instar larvae which can be used for all three genera of mosquitoes. A standard FISH protocol 12 is optimized for using BAC clones of genomic DNA as a probe on mitotic chromosomes of Ae. aegypti and Cx. quinquefasciatus, and for utilizing an intergenic spacer (IGS) region of ribosomal DNA (rDNA) as a probe on An. gambiae chromosomes. In addition to physical mapping, the developed technique can be applied to population cytogenetics and chromosome taxonomy/systematics of mosquitoes and other insect groups.

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Among the three mosquito genera, namely Anopheles, Aedes, and Culex, physical genome mapping techniques were established only for Anopheles, whose members possess readable polytene chromosomes. For the genera of Aedes and Culex, however, cytogenetic mapping remains challenging because of the poor quality of polytene chromosomes. Here we present a universal protocol for obtaining high-quality preparations of mitotic chromosomes and an optimized FISH protocol for all three genera of mosquitoes.

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. ...

Use of a Recombinant Mosquito Densovirus As a Gene Delivery Vector for the Functional Analysis of Genes in Mosquito Larvae

In vivo microinjection is the most commonly used gene transfer technique for analyzing the gene functions in individual mosquitoes. However, this method requires a more technically demanding operation and involves complicated procedures, especially when used in larvae due to their small size, relatively thin and fragile cuticle, and high mortality, which limit its application. In contrast, viral vectors for gene delivery have been developed to surmount extracellular and intracellular barriers. These systems have the advantages of easy manipulation, high gene transduction efficiency, long-term maintenance of gene expression, and the ability to produce persistent effects in vivo. Mosquito densoviruses (MDVs) are mosquito-specific, small single-stranded DNA viruses that can effectively deliver foreign nucleic acids into mosquito cells; however, the replacement or insertion of foreign genes to create recombinant viruses typically causes a loss of packaging and/or replication abilities, which is a barrier to the development of these viruses as delivery vectors.
Herein, we report using an artificial intronic small-RNA expression strategy to develop a non-defective recombinant Aedes aegypti densovirus (AaeDV) in vivo delivery system. Detailed procedures for the construction, packaging and quantitative analysis of the rAaeDV vectors, and for larval infection are described.
This study demonstrates, for the first time, the feasibility of developing a non-defective recombinant MDV micro RNA (miRNA) expression system, and thus providing a powerful tool for the functional analysis of genes in mosquito and establishing a basis for the application of viral paratransgenesis for controlling mosquito-borne diseases. We demonstrated that Aedes albopictus 1st instar larvae could be easily and effectively infected by introducing the virus into the water body of the larvae breeding site and that the developed rAaeDVs could be used to overexpress or knock down the expression of a specific target gene in larvae, providing a tool for the functional analysis of mosquito genes.

We report using an artificial intronic small RNA expression strategy to develop a non-defective recombinant Aedes aegypti densovirus (AaeDV) in vivo delivery system. A detailed procedure for the construction, packaging, and quantitative analysis of the rAaeDV vectors as well as for larval infection is described.

In vivo microinjection is the most commonly used gene transfer technique for analyzing the gene functions in individual mosquitoes. However, this method ...

Developmental Biology

Small-Cage Laboratory Trials of Genetically-Engineered Anopheline Mosquitoes

Control of mosquito-borne pathogens using genetically-modified vectors has been proposed as a promising tool to complement conventional control strategies. CRISPR-based homing gene drive systems have made transgenic technologies more accessible within the scientific community. Evaluation of transgenic mosquito performance and comparisons with wild-type counterparts in small laboratory cage trials provide valuable data for the design of subsequent field cage experiments and experimental assessments to refine the strategies for disease prevention. Here, we present three different protocols used in laboratory settings to evaluate transgene spread in anopheline mosquito vectors of malaria. These include inundative releases (no gene-drive system), and gene-drive overlapping and non-overlapping generation trials. The three trials vary in a number of parameters and can be adapted to desired experimental settings. Moreover, insectary studies in small cages are part of the progressive transition of engineered insects from the laboratory to open field releases. Therefore, the protocols described here represent invaluable tools to provide empirical values that will ultimately aid field implementation of new technologies for malaria elimination.

The protocols reported here illustrate three alternative ways to assess the performance of genetically-engineered mosquitoes destined for vector control in laboratory-contained small cage trials. Each protocol is tailored to the specific modification the mosquito strain bears (gene drive or non-gene drive) and the types of parameters measured.

Control of mosquito-borne pathogens using genetically-modified vectors has been proposed as a promising tool to complement conventional control ...

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

Mosquito gene editing has become routine in several laboratories with the establishment of systems such as transcription-activator-like effector nucleases (TALENs), zinc-finger nucleases (ZFNs), and homing endonucleases (HEs). More recently, clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) technology has offered an easier and cheaper alternative for precision genome engineering. Following nuclease action, DNA repair pathways will fix the broken DNA ends, often introducing indels. These out-of-frame mutations are then used for understanding gene function in the target organisms. A drawback, however, is that mutant individuals carry no dominant marker, making identification and tracking of mutant alleles challenging, especially at scales needed for many experiments. 
High-resolution melt analysis (HRMA) is a simple method to identify variations in nucleic acid sequences and utilizes PCR melting curves to detect such variations. This post-PCR analysis method uses fluorescent double-stranded DNA-binding dyes with instrumentation that has temperature ramp control data capture capability and is easily scaled to 96-well plate formats. Described here is a simple workflow using HRMA for the rapid detection of CRISPR/Cas9-induced indels and the establishment of mutant lines in the mosquito Ae. aegypti. Critically, all steps can be performed with a small amount of leg tissue and do not require sacrificing the organism, allowing genetic crosses or phenotyping assays to be performed after genotyping.

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

This article details a protocol for rapid identification of indels induced by CRISPR/Cas9 and selection of mutant lines in the mosquito Aedes aegypti using high-resolution melt analysis.

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

A Multi-detection Assay for Malaria Transmitting Mosquitoes

The Anopheles gambiae species complex includes the major malaria transmitting mosquitoes in Africa. Because these species are of such medical importance, several traits are typically characterized using molecular assays to aid in epidemiological studies. These traits include species identification, insecticide resistance, parasite infection status, and host preference. Since populations of the Anopheles gambiae complex are morphologically indistinguishable, a polymerase chain reaction (PCR) is traditionally used to identify species. Once the species is known, several downstream assays are routinely performed to elucidate further characteristics. For instance, mutations known as KDR in a para gene confer resistance against DDT and pyrethroid insecticides. Additionally, enzyme-linked immunosorbent assays (ELISAs) or Plasmodium parasite DNA detection PCR assays are used to detect parasites present in mosquito tissues. Lastly, a combination of PCR and restriction enzyme digests can be used to elucidate host preference (e.g., human vs. animal blood) by screening the mosquito bloodmeal for host-specific DNA. We have developed a multi-detection assay (MDA) that combines all of the aforementioned assays into a single multiplex reaction genotyping 33SNPs for 96 or 384 samples at a time. Because the MDA includes multiple markers for species, Plasmodium detection, and host blood identification, the likelihood of generating false positives or negatives is greatly reduced from previous assays that include only one marker per trait. This robust and simple assay can detect these key mosquito traits cost-effectively and in a fraction of the time of existing assays.

Malaria transmitting mosquitoes have a number of epidemiologically important characteristics that can only be detected using molecular techniques. Utilizing a MALDI-TOF based SNP genotyping platform, we developed an assay for simultaneously detecting multiple key traits (species, insecticide resistance, parasite infection and host choice) of malaria vectors.

The Anopheles gambiae species complex includes the major malaria transmitting mosquitoes in Africa. Because these species are of such medical importance, ...

Biology

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 ...

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;

Cyfrowa reakcja łańcuchowa polimerazy kropelkowej do wykrywania mutacji indel w genach docelowych

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

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

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Przeglądaj więcej filmów

Transfection and Mutagenesis of Target Genes in Mosquito Cells by Locked Nucleic Acid-modified Oligonucleotides

A Simple Chelex Protocol for DNA Extraction from Anopheles spp.

Multiplexed Isothermal Amplification Based Diagnostic Platform to Detect Zika, Chikungunya, and Dengue 1

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Use of a Recombinant Mosquito Densovirus As a Gene Delivery Vector for the Functional Analysis of Genes in Mosquito Larvae

Small-Cage Laboratory Trials of Genetically-Engineered Anopheline Mosquitoes

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

A Multi-detection Assay for Malaria Transmitting Mosquitoes

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

Cyfrowa reakcja łańcuchowa polimerazy kropelkowej do wykrywania mutacji indel w genach docelowych