This work was supported by Shenzhen Science and Technology Program (Grant No. KQTD20180411143628272) and special funds for science technology innovation and industrial development of Shenzhen Dapeng New District (Grant No. PT202101-02).

1. Target design and in vitro synthesis of sgRNA
<ol>
	<li>Predict the structure of target genes of interest and determine the boundaries between exons and introns via bioinformatic analysis of the B. dorsalis genome (software applications used here are listed in the Table of Materials). 
	NOTE: BLAT20 was used to search potential gene loci in the genome. The high-quality RNA-seq reads (transcriptome) were aligned to the acquired gene loci using Hisat221. Samtools22 was used to generate the sorted bam files. The sorted bam files were input to Stringtie223 to provide the assemble transcripts. The assemble transcripts and gene loci information were combined by Transdecoder24. The results acquired from Transdecoder were visualized in IGV tools25 and the boundaries between exons and introns could be determined.</li>
	<li>Identify the suitable target regions within the candidate target gene site. The total length must be less than 750 bp for more convenient sequencing (Figure 1B). Design specific primers to amplify the target area from wild-type genomic DNA by PCR (Figure 1B) (Primers: F-primer: AACATTGAATATCTGGAATCAGGTAAACT, R-primer: CCTCATTGTTGATTAATTCCGACTTC). Clone the PCR products into a blunt end-vector26 and select 20 individual bacterial colonies for sequencing to determine how conserved the target region is in the laboratory insect populations.</li>
	<li>A typical target site contains a three-nucleotide sequence motif (NGG or CCN) and a 20 bp sequence adjacent to the NGG or CCN motifs (20 bp-NGG or CCN-20 bp). Blast the candidate target sites against the B. dorsalis genome and make sure the predicted efficiency is high enough and the off-targeting rate is low; several open-source software programs can automatically predict this. In this protocol, sgRNAcas9 -AI27 is used to select and evaluate the optimal targets. Details of use can be found in the manual for this software. 
	NOTE: Possible target sites closed to the 5&#39; UTR of the target gene and 1-2 Gs at the start of the 20 bp sequence are favored. In order to achieve a large deletion which will be identified by PCR followed by agarose gel electrophoresis, designing two targets28 separated by over 100 bp is recommended (Figure 1C).</li>
	<li>Use the commercially available gRNA synthesis kit to generate the designed sgRNA. Perform each step following the user&#39;s guide. Resuspend the gRNA product in nuclease-free water, quantify the concentration using a UV-Vis spectrophotometer, and store at -80 &#176;C prior to use (the concentration of successfully synthesized single gRNA [10 &#181;L] with the kit used here is about 4000-6000 ng/&#181;L, or even higher). 
	&#8203;NOTE: Although generating the mutant flies is the first step for every researcher, the appropriate negative control for downstream experiment/analysis is critical. Generating these controls alongside the mutants saves researchers time and effort. For example, set scrambled sgRNA as a negative control.</li>
</ol>
2. Embryo collection and preparation
<ol>
	<li>Place B. dorsalis pupae&#160;into plastic cages. Provide a mixture of sugar and yeast (1:1) as food along with a water source after the adults eclose (Figure 2A, B). The rearing conditions are 55% relative humidity (RH), 26.5 &#176;C, and a 14:10 L/D cycle(lights on at six in the morning, lights off at eight in the evening).</li>
	<li>Most adults reach sexual maturity 10 days after emergence. Provide a suitable environment to help adult flies mate as much as possible. Ideally, use a light stand with a light of 30-50 lux. This can improve the fecundity of females, therefore, improving the efficiency of embryo collection. 
	NOTE: Putting the adults (aged 5-6 days after emergence) in a dimly lit (&#60;100 lux) environment can promote mating. Generally, the peak of oviposition happens at 03:00 p.m. (14:10/L:D, lights on 06:00 a.m., lights off at 08:00 p.m.); to obtain enough embryos, placing oviposition chambers in the cages 30 min before 03:00 p.m. is recommended.</li>
	<li>Place a 200-mesh gauze in the oviposition chamber, 1-2 mm away from the chamber lid. This will help with obtaining as many embryos as possible. 
	NOTE: Avoid letting the embryos get soaked in orange juice or rubbed with gauze, as this can significantly decrease the embryos&#39; survival rate.</li>
	<li>Put a new oviposition chamber into the cage when the microinjection setup is ready. Collect the embryos every 10 min using a fine wet brush, then line them up on a self-made injection plate (plexiglass with a length of 55 mm, a width of 13.75 mm, and a height of 5 mm, A 45 mm x 5 mm x 0.3 mm shallow groove is opened in the middle to facilitate egg placement). If the embryos have high internal pressure, slight desiccation at &#60;10% RH for 10 min is optional. Dip the embryos into halocarbon oil during injection to avoid further desiccation (Figure 2C).</li>
</ol>
3. Microinjection of the embryo
<ol>
	<li>Prepare the glass injection needle using a micropipette puller. 
	NOTE: Setting the parameters following the user guide is important. In these protocols, different needle shapes can be made using the parameters suggested by the manual of the micropipette puller. The glass capillary must be as clean as possible to prevent dust from clogging the needle.</li>
	<li>Prepare the working solution. Mix the Cas9 protein and the corresponding sgRNA to the following working concentrations:&#160;sgRNA, 300 ng/&#181;L; Cas9 protein, 150 ng/&#181;L. Add 1 &#181;L of phenol red to the mixture to serve as a convenient way to mark injected embryos. Place the prepared mixture on ice to avoid degradation of the sgRNA. 
	NOTE: Use nuclease-free pipette tips and PCR tubes. The concentration of Cas9 protein needs to be less than or equal to 150 ng/&#181;L; higher concentrations are toxic and significantly decrease the embryo survival rate. Cas9 could also be delivered in DNA or mRNA format to achieve successful gene editing17,19. In this protocol, Cas9 protein is recommended since mRNAs are susceptible to degradation, and the expression of Cas9 protein from plasmid DNA takes time for transcription and translation.</li>
	<li>Set the parameters for the injector. The initial program is Pi-500 hPa, Ti-0.5 s, and PC-200 hPa. Adjust these parameters further as needed during microinjection.</li>
	<li>Add 3 &#181;L of the mixture into the injection needle. Avoid introducing air bubbles that can possibly clog the needle. Open the needle using a Microgrinder.</li>
	<li>Connect the needle to the micromanipulator according to the user guide (Figure 2E).</li>
	<li>Put the plate with lined-up embryos on the objective table. Adjust the position of the micropipette under a fine optical microscope, setting the micropipette and embryo on the same plane. Adjust the droplet volume by pressing the pedal; 1/10 the volume of embryos is recommended.</li>
	<li>Insert the needle tip into the posterior (vegetal pole) of the embryo. Deliver the mixtures into the embryo by pressing the pedal from the injector. If phenol red is added, a slightly reddish color is observed instantly. 
	&#8203;NOTE: A small volume of cytoplasmic backflow from the pinhole will not decrease the survival rate of the embryo. Injection of 200 embryos is enough for successful mutagenesis of one gene. Adding more halocarbon oil to immerse the embryos can prevent further desiccation. The injection needs to be performed before pole cell formation, which ensures every mutation can be efficiently inherited.</li>
</ol>
4. Post-injection insect rearing
<ol>
	<li>Put the injection plate with the injected embryos into an artificial climate chamber kept at 55% RH, 26.5 &#176;C, and a 14:10 L/D cycle(lights on at six in the morning, lights off at eight in the evening).</li>
	<li>After injection, the embryo formation usually takes 24 h to complete and 36 h to hatch. Use a fine-tip brush to transfer the larvae onto a prepared larval diet. Collect larvae three or four times daily until no more larvae hatch. Generally, larvae take 2-3 days to finish hatching and pupate within 7 days. 
	&#8203;NOTE: A maize-based diet was used to feed the larvae. The recipe contains 150 g of corn flour, 150 g of banana, 0.6 g of sodium benzoate, 30 g of yeast, 30 g of sucrose, 30 g of paper towel, 1.2 mL of hydrochloric acid, and 300 mL of water29. Minimize the time larvae are left in oil (&#60;2 h). Move the larvae onto the diet immediately after hatching as far as possible. This can significantly increase the survival and pupation rates. Do not provide too much diet (a diet with 2/3 of a 90 mm Petri dish is enough for 30 larvae); otherwise, it will become moldy and decrease the larval survival rate.</li>
	<li>Put the mature larvae into the wet sand to pupate. The pupation stage takes about 10 days. Move the pupae to plastic cages before eclosion begins.</li>
</ol>
5. Mutant screening
<ol>
	<li>A flow chart of the mutant screening is illustrated in Figure 3A. Cross G0 adults obtained by microinjection with wild-type adults to obtain heterozygous lines. Verify the type of mutations the offspring (G1) carry by genotyping.</li>
	<li>Self-cross G1 with the same mutant genotype to obtain homozygous lines in G2. If genotypes of the mutants in G1 are entirely different, cross the heterozygotes with wild-type flies. Homozygous lines are usually obtained in G3. Maintain two or three homozygous lines for subsequent phenotyping experiments.</li>
	<li>Use the genomic DNA from a fresh puparium or a single mid-leg of individual adults to perform genotyping. Design specific primers to amplify the target area; the primers must set at least 50 bps upstream and downstream of the target site. Refer to step 1.2 for the primer details. 
	NOTE: Since the amount of the DNA in a puparium is extremely low, commercially available DNA extraction kits are recommended.</li>
	<li>Perform PCR using the following cycling conditions: 98 &#176;C for 3 min, followed by 35 cycles of 98 &#176;C for 10 s, 15 s at an annealing temperature appropriate for the designed primers, 72 &#176;C for 35 s, followed by a final extension of 72 &#176;C for 10 min.</li>
	<li>Sequence the purified PCR products. When multiple overlapping peaks adjacent to the target site are detected (Figure 3B), successful mutagenesis has occurred. Sub-clone the purified PCR products into a blunt-end vector and select 10 individual bacterial colonies for sequencing to verify the genotype of the mutants. Maintain the lines with frameshift mutations and premature translation terminations (Figure 3C). 
	NOTE: There is no need to perform single clone sequencing to verify the genotype of G0 individuals. Just sequence the PCR products and maintain the individuals with obvious multiple overlap peaks adjacent to the target site. After the initial injection, selecting 10 embryos to extract genomic DNA and sequencing the PCR products based on the standards in step 5.3 is recommended. This can help to predict mutation rates in advance. In this study, sanger sequencing to determine the genotype was done by a sequencing company.</li>
</ol>

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H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Knockout+of+juvenile+hormone+receptor,+Methoprene-tolerant,+induces+black+larval+phenotype+in+the+yellow+fever+mosquito,+Aedes+aegypti.">Knockout of juvenile hormone receptor, Methoprene-tolerant, induces black larval phenotype in the yellow fever mosquito, Aedes aegypti.</a> Proceedings of the National Academy of Sciences. 116 (43), 21501-21507 (2019).</li><li>Laohakieat, K., Isasawin, S., Thanaphum, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+transformer-2+and+fruitless+characterisation+with+developmental+expression+profiles+of+sex-determining+genes+in+Bactrocera+dorsalis+and+B.+correcta.">The transformer-2 and fruitless characterisation with developmental expression profiles of sex-determining genes in Bactrocera dorsalis and B. correcta.</a> Scientific Reports. 10 (1), 1-13 (2020).</li><li>Gabrieli, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sperm-less+males+modulate+female+behaviour+in+Ceratitis+capitata+(Diptera:+Tephritidae).">Sperm-less males modulate female behaviour in Ceratitis capitata (Diptera: Tephritidae).</a> Insect Biochemistry and Molecular Biology. 79, 13-26 (2016).</li></ol>

The authors do not have any conflicts of interest.

The CRISPR/Cas9 system is the most widely used gene editing tool and has various applications, such as gene threpy30, crop breeding31, and basic studies of gene fuctions32. This system has already been applied for gene editing in various insect species and has served as an effective tool for functional gene studies in pests. The protocols we present here standardize the procedure of design and synthesis of guide RNAs, collecting embryos, embryo injec...

The Oriental fruit fly, Bactrocera dorsalis, is a cosmopolitan insect pest species that causes damage to over 150 species of fruit crops, including guava, mango, Eugenia spp., Surinam cherry, citrus, loquat, and papaya1. The damage caused in Guangdong Province (China) alone is estimated at over 200 million yuans. Adult females insert their eggs beneath the skin of ripening or ripened fruits, causing decay and abscission of the fruit, which decreases fruit quality and overall yield of the crop2. Since adult fruit flies have great flight capacity and their larvae bore into the fruit skin, insecticides requiring direct contact with the pest perform poorly in the field. Additionally, the extensive use of insecticides has increased the resistance of B. dorsalis against various agricultural chemicals, making control of these damaging pests even more difficult3. Therefore, the development of effective and environmentally friendly pest management strategies is desperately needed.
Recently, with the development of molecular biological tools and high-throughput sequencing technologies, scientists are attempting to develop environmentally-friendly pest management strategies, such as RNAi, that target the functionality of important genes (molecular targets) of various insect pests. Genes that are critical to the survival and reproduction of the pest can be identified through functional gene studies and further serve as potential molecular targets for the improvement of specifically targeted and environmentally friendly pest management tools4. To adapt such strategies to Oriental fruit fly control, effective methods for functional gene research are needed.
The CRISPR/Cas (clustered regularly interspaced short palindromic repeats/CRISPR-associated) endonuclease system was initially discovered in bacteria and archaea and found to be an adaptive mechanism involved in the recognition and degradation of foreign intracellular DNA, such as that introduced by infecting bacteriophages5. In the type II CRISPR system, Cas9 endonuclease is guided by small associated RNAs (crRNA and tracrRNA) to cleave trespassing DNA6,7,8 and has become one of the most widely used tools for gene-editing to date9,10,11,12. Since the CRISPR/Cas9 system has several advantages, such as high efficiency of gene silencing and low cost, it has already been applied for gene editing in various insect species, including Aedes aegypti13,14, Locusta migratoria15, and Bombyx mori16. In B. dorsalis, genes related to body color, wing dimorphism, and sex determination have been successfully knocked out using CRISPR/Cas917,18,19. However, detailed procedures for CRISPR/Cas9 application in this insect remain incomplete. Moreover, some steps provided by researchers for B. dorsalis gene editing are also varied and in need of standardization. For example, the forms of Cas9 were different in published references17,18,19.
This paper provides a systematic method for mutagenesis of B. dorsalis using the CRISPR/Cas9 system, including the design and synthesis of guide RNAs, collecting embryos, embryo injection, insect rearing, and mutant screening. This protocol will serve as a useful guide for generating mutant flies for researchers who are interested in the functional gene studies in B. dorsalis.

<table><tbody><tr><td>6x DNA Loading Buffer</td><td>TransGen Biotech</td><td>GH101-01</td><td></td></tr><tr><td>Artificial climate chamber</td><td>ShangHai BluePard</td><td>MGC-350P</td><td></td></tr><tr><td>AxyPrep Genomic DNA Mini-Extraction Kit</td><td>Axygen</td><td>AP-MN-MS-GDNA-250G</td><td></td></tr><tr><td>BLAT</td><td>NA</td><td>NA</td><td>For searching potential gene loci in the genome</td></tr><tr><td>Capillary Glass</td><td>WPI&amp;nbsp;</td><td>1B100F-4</td><td></td></tr><tr><td>Eppendorf InjectMan 4 micromanipulator</td><td>Eppendorf</td><td>InjectMan 4</td><td></td></tr><tr><td>GeneArt Precision gRNA Synthesis Kit</td><td>Thermo Fisher Scientific</td><td>A29377</td><td></td></tr><tr><td>Hisat2</td><td>NA</td><td>NA</td><td>For aligning the transcriptome to the acquired gene loci</td></tr><tr><td>IGV</td><td>NA</td><td>NA</td><td>For visualizing the results from Transdecoder</td></tr><tr><td>Microgrinder</td><td>NARISHIGE</td><td>EG-401</td><td></td></tr><tr><td>Olympus Microscope</td><td>Olympus Corporation</td><td>SZ2-ILST</td><td></td></tr><tr><td>pEASY-Blunt Cloning Kit</td><td>TransGen Biotech</td><td>CB101-02</td><td>https://www.transgenbiotech.com/data/upload/pdf/CB101_2022-07-14.pdf</td></tr><tr><td>Phenol red solution</td><td>Sigma-Aldrich</td><td>P0290-100ML</td><td></td></tr><tr><td>Pipette cookbook 2018 P-97 &amp;amp; P-1000 Micropipette Pullers</td><td>Instrument Company&amp;nbsp;</td><td></td><td>https://www.sutter.com/PDFs/cookbook.pdf</td></tr><tr><td>PrimeSTAR HS (Premix)</td><td>Takara Biomedical Technology</td><td>R040A</td><td></td></tr><tr><td>SAMtools</td><td>NA</td><td>NA</td><td>For generating the sorted bam files</td></tr><tr><td>sgRNAcas9-AI</td><td>NA</td><td>NA</td><td>sgRNA design&lt;br/&gt; http://123.57.239.141:8080/home</td></tr><tr><td>Sutter Micropipette Puller Sutter&amp;nbsp;</td><td>Instrument Company&amp;nbsp;</td><td>P-97</td><td></td></tr><tr><td>Trans2K DNA Marker</td><td>TransGen Biotech</td><td>BM101-02</td><td></td></tr><tr><td>Transdecoder</td><td>NA</td><td>NA</td><td>For combining the results of assemble transcripts and gene loci information&lt;br/&gt; https://github.com/TransDecoder/TransDecoder/releases/tag/TransDecoder-v5.5.0</td></tr><tr><td>TrueCut Cas9 Protein v2</td><td>Thermo Fisher Scientific</td><td>A36498</td><td></td></tr><tr><td>Ultra-trace biological detector</td><td>Thermo Fisher Scientific</td><td>Nanodrop 2000C</td><td></td></tr></tbody></table>

protocols for crispr/cas9 mutagenesis of the oriental fruit fly bactrocera dorsalis

The Oriental fruit fly, Bactrocera dorsalis, is a highly invasive and adaptive pest species that causes damage to citrus and over 150 other fruit crops worldwide. Since adult fruit flies have great flight capacity and females lay their eggs under the skins of fruit, insecticides requiring direct contact with the pest usually perform poorly in the field. With the development of molecular biological tools and high-throughput sequencing technology, many scientists are attempting to develop environmentally friendly pest management strategies. These include RNAi or gene editing-based pesticides that downregulate or silence genes (molecular targets), such as olfactory genes involved in searching behavior, in various insect pests. To adapt these strategies for Oriental fruit fly control, effective methods for functional gene research are needed. Genes with critical functions in the survival and reproduction of B. dorsalis serve as good molecular targets for gene knockdown and/or silencing. The CRISPR/Cas9 system is a reliable technique used for gene editing, especially in insects. This paper presents a systematic method for CRISPR/Cas9 mutagenesis of B. dorsalis, including the design and synthesis of guide RNAs, collecting embryos, embryo injection, insect rearing, and mutant screening. These protocols will serve as a useful guide for generating mutant flies for researchers interested in functional gene studies in B. dorsalis.

This protocol presents detailed steps for the development of B. dorsalis mutants using CRISPR/Cas9 technology, including representative results from gDNA selection, collecting embryos and microinjection, insect maintenance, and mutant screening.
The example of the target site of the selected gene is located in the third exon (Figure 1C). This site is highly conserved, and a single band was detected by gel electrophoresis for the DNA template for synthetic...

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Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis

This paper presents the step-by-step protocols for CRISPR/Cas9 mutagenesis of the Oriental fruit fly Bactrocera dorsalis. Detailed steps provided by this standardized protocol will serve as a useful guide for generating mutant flies for functional gene studies in B. dorsalis.

Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis

The Oriental fruit fly, Bactrocera dorsalis, is a highly invasive and adaptive pest species that causes damage to citrus and over 150 other fruit crops ...

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Biology

2.2K Views.  Northeast Forestry University. De oosterse fruitvlieg is een zeer invasieve en adaptieve plaagsoort. Effectieve methoden voor functioneel genonderzoek kunnen ons helpen om milieuvriendelijke strategieën voor ongediertebestrijding te ontwikkelen. Deze methode is hoog rendement en lage kosten.Ons protocol zal dienen als een nuttige gids voor het genereren van gemuteerde vliegen voor oosterse fruitvliegonderzoekers. Om te beginnen, voorspel de structuur van doelgenen van belang en bepaal de grenzen tussen exonen en in...

Video: Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide RNA (sgRNA) that confers specificity, the Cas9 protein cleaves both DNA strands at the targeted locus. The DNA break can trigger either non-homologous end joining (NHEJ) or homology directed repair (HDR). NHEJ can introduce small deletions or insertions which lead to frame-shift mutations, while HDR allows for larger and more precise perturbations. Here, we present protocols for generating knockout cell lines by coupling established CRISPR/Cas9 methods with two options for downstream selection/screening. The NHEJ approach uses a single sgRNA cut site and selection-independent screening, where protein production is assessed by dot immunoblot in a high-throughput manner. The HDR approach uses two sgRNA cut sites that span the gene of interest. Together with a provided HDR template, this method can achieve deletion of tens of kb, aided by the inserted selectable resistance marker. The appropriate applications and advantages of each method are discussed.

Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide ...

Genetics

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 ...

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

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

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

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

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

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been harnessed by researchers to study important biologically relevant problems. The unparalleled power of the CRISPR/Cas genome editing method allows researchers to precisely edit any locus of their choosing, thereby facilitating an increased understanding of gene function. Several methods for editing the C. elegans genome by CRISPR/Cas9 have been described previously. Here, we discuss and demonstrate a method which utilizes in vitro assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. Specifically, in this article, we go through the step-by-step process of introducing premature stop codons into the C. elegans rbm-3.2 gene by homology-directed repair using this method of CRISPR/Cas9 editing. This relatively simple editing method can be used to study the function of any gene of interest and allows for the generation of homozygous-edited C. elegans by CRISPR/Cas9 editing&#160;in less than two weeks.

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

The goal of this protocol is to enable seamless CRISPR/Cas9 editing of the C. elegans genome using assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. This protocol can be used to make a variety of genetic modifications in C. elegans including insertions, deletions, gene replacements and codon substitutions.

The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been ...

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 ...

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

Microinjection of Western Corn Rootworm, Diabrotica virgifera virgifera, Embryos for Germline Transformation, or CRISPR/Cas9 Genome Editing

The western corn rootworm (WCR) is an important pest of corn and is well known for its ability to rapidly adapt to pest management strategies. Although RNA interference (RNAi) has proved to be a powerful tool for studying WCR biology, it has its limitations. Specifically, RNAi itself is transient (i.e. does not result in long-term Mendelian inheritance of the associated phenotype), and it requires knowing the DNA sequence of the target gene. The latter can be limiting if the phenotype of interest is controlled by poorly conserved, or even novel genes, because identifying useful targets would be challenging, if not impossible. Therefore, the number of tools in WCR&#39;s genomic toolbox should be expanded by the development of methods that could be used to create stable mutant strains and enable sequence-independent surveys of the WCR genome. Herein, we detail the methods used to collect and microinject precellular WCR embryos with nucleic acids. While the protocols described herein are aimed at the creation of transgenic WCR, CRISPR/Cas9-genome editing could also be performed using the same protocols, with the only difference being the composition of the solution injected into the embryos.

Microinjection of Western Corn Rootworm, Diabrotica virgifera virgifera, Embryos for Germline Transformation, or CRISPR/Cas9 Genome Editing

Here we present protocols for collecting and microinjecting precellular western corn rootworm embryos for the purpose of performing functional-genomic assays such as germline transformation and CRISPR/Cas9-genome editing.

The western corn rootworm (WCR) is an important pest of corn and is well known for its ability to rapidly adapt to pest management strategies. Although ...

Bioengineering

Generation of Genomic Deletions in Mammalian Cell Lines via CRISPR/Cas9

The prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system may be re-purposed for site-specific eukaryotic genome engineering. CRISPR/Cas9 is an inexpensive, facile, and efficient genome editing tool that allows genetic perturbation of genes and genetic elements. Here we present a simple methodology for CRISPR design, cloning, and delivery for the production of genomic deletions. In addition, we describe techniques for deletion, identification, and characterization. This strategy relies on cellular delivery of a pair of chimeric single guide RNAs (sgRNAs) to create two double strand breaks (DSBs) at a locus in order to delete the intervening DNA segment by non-homologous end joining (NHEJ) repair. Deletions have potential advantages as compared to single-site small indels given the efficiency of biallelic modification, ease of rapid identification by PCR, predictability of loss-of-function, and utility for the study of non-coding elements. This approach can be used for efficient loss-of-function studies of genes and genetic elements in mammalian cell lines.

CRISPR/Cas9 is a robust system to produce disruption of genes and genetic elements. Here we describe a protocol for the efficient creation of genomic deletions in mammalian cell lines using CRISPR/Cas9.

The prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system may be re-purposed for site-specific ...

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 ...

Construction of Homozygous Mutants of Migratory Locust Using CRISPR/Cas9 Technology

The migratory locust, Locusta migratoria, is not only one of the worldwide plague locusts that caused huge economic losses to human beings but also an important research model for insect metamorphosis. The CRISPR/Cas9 system can accurately locate at a specific DNA locus and cleave within the target site, efficiently introducing double-strand breaks to induce target gene knockout or integrate new gene fragments into the specific locus. CRISPR/Cas9-mediated genome editing is a powerful tool for addressing questions encountered in locust research as well as a promising technology for locust control. This study provides a systematic protocol for CRISPR/Cas9-mediated gene knockout with the complex of Cas9 protein and single guide RNAs (sgRNAs) in migratory locusts. The selection of target sites and design of sgRNA are described in detail, followed by in vitro synthesis and verification of the sgRNAs. Subsequent procedures include egg raft collection and tanned-egg separation to achieve successful microinjection with low mortality rate, egg culture,&#160;preliminary estimation of the mutation rate, locust breeding as well as detection, preservation, and passage of the mutants to ensure population stability of the edited locusts. This method can be used as a reference for CRISPR/Cas9 based gene editing applications in migratory locusts as well as in other insects.

This study provides a systematically optimized procedure of CRISPR/Cas9 ribonuclease-based construction of homozygous locust mutants as well as a detailed method for cryopreservation and resuscitation of the locust eggs.

The migratory locust, Locusta migratoria, is not only one of the worldwide plague locusts that caused huge economic losses to human beings but also an ...

Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis

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