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.

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			<td>GH101-01</td>
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			<td>Artificial climate chamber</td>
			<td>ShangHai BluePard</td>
			<td>MGC-350P</td>
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			<td>AxyPrep Genomic DNA Mini-Extraction Kit</td>
			<td>Axygen</td>
			<td>AP-MN-MS-GDNA-250G</td>
			<td></td>
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		<tr>
			<td>BLAT</td>
			<td>NA</td>
			<td>NA</td>
			<td>For searching potential gene loci in the genome</td>
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			<td>Capillary Glass</td>
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			<td>Eppendorf</td>
			<td>InjectMan 4</td>
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			<td>GeneArt Precision gRNA Synthesis Kit</td>
			<td>Thermo Fisher Scientific</td>
			<td>A29377</td>
			<td></td>
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		<tr>
			<td>Hisat2</td>
			<td>NA</td>
			<td>NA</td>
			<td>For aligning the transcriptome to the acquired gene loci</td>
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			<td>IGV</td>
			<td>NA</td>
			<td>NA</td>
			<td>For visualizing the results from Transdecoder</td>
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			<td>Microgrinder</td>
			<td>NARISHIGE</td>
			<td>EG-401</td>
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			<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>
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		<tr>
			<td>Pipette cookbook 2018 P-97 &#38; P-1000 Micropipette Pullers</td>
			<td>Instrument Company&#160;</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 
			http://123.57.239.141:8080/home</td>
		</tr>
		<tr>
			<td>Sutter Micropipette Puller Sutter&#160;</td>
			<td>Instrument Company&#160;</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 
			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...

Watch this Scientific Journal Video about Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis at JoVE.com

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 ...

protocols-for-crisprcas9-mutagenesis-oriental-fruit-fly-bactrocera

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2.2K Views.  Northeast Forestry University. 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.

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

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular signatures, or to evolve proteins in the laboratory. Here, we present a protocol that allows the generation of large (>1011) mutant libraries for a given target sequence. This method is based on replication of a ColE1 plasmid encoding the desired sequence by a low-fidelity variant of DNA polymerase I (LF-Pol I). The target plasmid is transformed into a mutator strain of E. coli and plated on solid media, yielding between 0.2 and 1 mutations/kb, depending on the location of the target gene. Higher mutation frequencies are achieved by iterating this process of mutagenesis. Compared to alternative methods of mutagenesis, our protocol stands out for its simplicity, as no cloning or PCR are involved. Thus, our method is ideal for mutational labeling of plasmids or other Pol I templates or to explore large sections of sequence space for the evolution of activities not present in the original target. The tight spatial control that PCR or randomized oligonucleotide-based methods offer can also be achieved through subsequent cloning of specific sections of the library. Here we provide protocols showing how to create a random mutant library and how to establish drug-based selections in E. coli to identify mutants exhibiting new biochemical activities.

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

Here we demonstrate a simple protocol to create a random mutant library for a given target sequence. We show how this method, which is performed in vivo in Escherichia coli, can be coupled with functional selections to evolve new enzymatic activities.

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular ...

Isolation and Biophysical Study of Fruit Cuticles

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and abiotic stresses and also regulates water flow from the external environment.1 A biopolyester (cutin) and long-chain fatty acids (waxes) form the principal structural framework of the cuticle; the functional integrity of the cuticular layer depends on the outer 'epicuticular' layer as well as the blend consisting of the cutin biopolymer and 'intracuticular' waxes.2 Herein, we describe a comprehensive protocol to extract waxes exhaustively from commercial tomato (Solanum lycopersicum) fruit cuticles or to remove epicuticular and intracuticular waxes sequentially and selectively from the cuticle composite. The method of Jetter and Sch&auml;ffer (2001) was adapted for the stepwise extraction of epicuticular and intracuticular waxes from the fruit cuticle.3,4 To monitor the process of sequential wax removal, solid-state cross-polarization magic-angle-spinning (CPMAS) 13C NMR spectroscopy was used in parallel with atomic force microscopy (AFM), providing molecular-level structural profiles of the bulk materials complemented by information on the microscale topography and roughness of the cuticular surfaces. To evaluate the cross-linking capabilities of dewaxed cuticles from cultivated wild-type and single-gene mutant tomato fruits, MAS 13C NMR was used to compare the relative proportions of oxygenated aliphatic (CHO and CH2O) chemical moieties.
Exhaustive dewaxing by stepwise Soxhlet extraction with a panel of solvents of varying polarity provides an effective means to isolate wax moieties based on the hydrophobic characteristics of their aliphatic and aromatic constituents, while preserving the chemical structure of the cutin biopolyester. The mechanical extraction of epicuticular waxes and selective removal of intracuticular waxes, when monitored by complementary physical methodologies, provides an unprecedented means to investigate the cuticle assembly: this approach reveals the supramolecular organization and structural integration of various types of waxes, the architecture of the cutin-wax matrix, and the chemical composition of each constituent. In addition, solid-state 13C NMR reveals differences in the relative numbers of CHO and CH2O chemical moieties for wild-type and mutant red ripe tomato fruits. The NMR techniques offer exceptional tools to fingerprint the molecular structure of cuticular materials that are insoluble, amorphous, and chemically heterogeneous. As a noninvasive surface-selective imaging technique, AFM furnishes an effective and direct means to probe the structural organization of the cuticular assembly on the nm-&mu;m length scale.

Aerial plant organs are protected by the cuticle, a supramolecular biopolyester-wax assembly. We present protocols to monitor selective removal of epi- and intracuticular waxes from tomato fruit cuticles on molecular and micro scales by solid-state NMR and atomic force microscopy, respectively, and to assess the cross-linking capacity of engineered cuticular biopolyesters.

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and ...

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 ...

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these independently derived phenotypes, which share a common genetic architecture. This is perhaps best exemplified by the numerous convergent features of gymnotiforms and mormyrids, two species-rich teleost clades that produce and detect weak electric fields and are called weakly electric fish. In the 50 years since the discovery that weakly electric fish use electricity to sense their surroundings and communicate, a growing community of scientists has gained tremendous insights into evolution of development, systems and circuits neuroscience, cellular physiology, ecology, evolutionary biology, and behavior. More recently, there has been a proliferation of genomic resources for electric fish. Use of these resources has already facilitated important insights with regards to the connection between genotype and phenotype in these species. A major obstacle to integrating genomics data with phenotypic data of weakly electric fish is a present lack of functional genomics tools. We report here a full protocol for performing CRISPR/Cas9 mutagenesis that utilizes endogenous DNA repair mechanisms in weakly electric fish. We demonstrate that this protocol is equally effective in both the mormyrid species Brienomyrus brachyistius and the gymnotiform Brachyhypopomus gauderio by using CRISPR/Cas9 to target indels and point mutations in the first exon of the sodium channel gene scn4aa. Using this protocol, embryos from both species were obtained and genotyped to confirm that the predicted mutations in the first exon of the sodium channel scn4aa were present. The knock-out success phenotype was confirmed with recordings showing reduced electric organ discharge amplitudes when compared to uninjected size-matched controls.

Here, a protocol is presented to produce and rear CRISPR/Cas9 genome knockout electric fish. Outlined in detail are the required molecular biology, breeding, and husbandry requirements for both a gymnotiform and a mormyrid, and injection techniques to produce Cas9-induced indel F0 larvae.

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these ...

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 ...

Genome Engineering of Primary Human B Cells Using CRISPR/Cas9

B cells are lymphocytes derived from hematopoietic stem cells and are a key component of the humoral arm of the adaptive immune system. They make attractive candidates for cell-based therapies because of their ease of isolation from peripheral blood, their ability to expand in vitro, and their longevity in vivo. Additionally, their normal biological function&#8212;to produce large amounts of antibodies&#8212;can be utilized to express very large amounts of a therapeutic protein, such as a recombinant antibody to fight infection, or an enzyme for the treatment of enzymopathies. Here, we provide detailed methods for isolating primary human B cells from peripheral blood mononuclear cells (PBMCs) and activating/expanding isolated B cells in vitro. We then demonstrate the steps involved in using the CRISPR/Cas9 system for site-specific KO of endogenous genes in B cells. This method allows for efficient KO of various genes, which can be used to study the biological functions of genes of interest. We then demonstrate the steps for using the CRISPR/Cas9 system together with a recombinant, adeno-associated, viral (rAAV) vector for efficient site-specific integration of a transgene expression cassette in B cells. Together, this protocol provides a step-by-step engineering platform that can be used in primary human B cells to study biological functions of genes as well as for the development of B-cell therapeutics.

Here we provide a detailed, step-by-step protocol for CRISPR/Cas9-based genome engineering of primary human B cells for gene knockout (KO) and knock-in (KI) to study biological functions of genes in B cells and the development of B-cell therapeutics.

B cells are lymphocytes derived from hematopoietic stem cells and are a key component of the humoral arm of the adaptive immune system. They make ...

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 ...

CRISPR/Cas9 Gene Editing of Hematopoietic Stem and Progenitor Cells for Gene Therapy Applications

CRISPR/Cas9 is a highly versatile and efficient gene-editing tool adopted widely to correct various genetic mutations. The feasibility of gene manipulation of hematopoietic stem and progenitor cells (HSPCs) in vitro makes HSPCs an ideal target cell for gene therapy. However, HSPCs moderately lose their engraftment and multilineage repopulation potential in ex vivo culture. In the present study, ideal culture conditions are described that improves HSPC engraftment and generate an increased number of gene-modified cells in vivo. The current report displays optimized in vitro culture conditions, including the type of culture media, unique small molecule cocktail supplementation, cytokine concentration, cell culture plates, and culture density. In addition to that, an optimized HSPC gene-editing procedure, along with the validation of the gene-editing events, are provided. For in vivo validation, the gene-edited HSPCs infusion and post-engraftment analysis in mouse recipients are displayed. The results demonstrated that the culture system increased the frequency of functional HSCs in vitro, resulting in robust engraftment of gene-edited cells in vivo.

The present protocol describes an optimized hematopoietic stem and progenitor cell (HSPC) culture procedure for the robust engraftment of gene-edited cells in vivo.

CRISPR/Cas9 is a highly versatile and efficient gene-editing tool adopted widely to correct various genetic mutations. The feasibility of gene ...

CRISPR-Cas9-Mediated Precise Knock-In Edits in Zebrafish Hearts

Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological phenomena. Zebrafish have been used as an effective genetic model to study numerous questions related to heritable disease, development, and toxicology at the whole-organ and -organism level. Due to the well-annotated and mapped zebrafish genome, numerous tools for gene editing have been developed. However, the efficacy of generating and ease of detecting precise knock-in edits using CRISPR is a limiting factor. Described here is a CRISPR-Cas9-based knock-in approach with the simple detection of precise edits in a gene responsible for cardiac repolarization and associated with the electrical disorder, Long QT Syndrome (LQTS). This two-single-guide RNA (sgRNA) approach excises and replaces the target sequence and links a genetically encoded reporter gene. The utility of this approach is demonstrated by describing non-invasive phenotypic measurements of cardiac electrical function in wild-type and gene-edited zebrafish larvae. This approach enables the efficient study of disease-associated variants in a whole organism. Furthermore, this strategy offers possibilities for the insertion of exogenous sequences of choice, such as reporter genes, orthologs, or gene editors.

This protocol describes an approach to facilitate precise knock-in edits in zebrafish embryos using CRISPR-Cas9 technology. A phenotyping pipeline is presented to demonstrate the applicability of these techniques to model a Long QT Syndrome-associated gene variant.

Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological ...

Centromere-Associated Protein-E Gene Editing Using the CRISPR/Cas9 System

Generation of Centromere-Associated Protein-E CENP-E-/- Knockout Cell Lines using the CRISPR/Cas9 System

The CRISPR (clustered regularly interspaced short palindromic repeats)/Cas9 system has emerged as a powerful tool for precise and efficient gene editing in a variety of organisms. Centromere-associated protein-E (CENP-E) is a plus-end-directed kinesin required for kinetochore-microtubule capture, chromosome alignment, and spindle assembly checkpoint. Although cellular functions of the CENP-E proteins have been well studied, it has been difficult to study the direct functions of CENP-E proteins using traditional protocols because CENP-E ablation usually leads to spindle assembly checkpoint activation, cell cycle arrest, and cell death. In this study, we have completely knocked out the CENP-E gene in human HeLa cells and successfully generated the CENP-E-/- HeLa cells using the CRISPR/Cas9 system.
Three optimized phenotype-based screening strategies were established, including cell colony screening, chromosome alignment phenotypes, and the fluorescent intensities of CENP-E proteins, which effectively improve the screening efficiency and experimental success rate of the CENP-E knockout cells. Importantly, CENP-E deletion results in chromosome misalignment, the abnormal location of the BUB1 mitotic checkpoint serine/threonine kinase B (BubR1) proteins, and mitotic defects. Furthermore, we have utilized the CENP-E knockout HeLa cell model to develop an identification method for CENP-E-specific inhibitors.
In this study, a useful approach to validate the specificity and toxicity of CENP-E inhibitors has been established. Moreover, this paper presents the protocols of CENP-E gene editing using the CRISPR/Cas9 system, which could be a powerful tool to investigate the mechanisms of CENP-E in cell division. Moreover, the CENP-E&#160;knockout cell line would contribute to the discovery and validation of CENP-E inhibitors, which have important implications for antitumor drug development, studies of cell division mechanisms in cell biology, and clinical applications.

Author Spotlight: Establishing CENP-E Knockout HeLa Cells &#8211; A Novel Approach to Study Kinesin-7 CENP-E Biology and its Inhibitors

This article reports the construction of centromere-associated protein-E (CENP-E) knockout cells using the CRISPR/Cas9 system and three phenotype-based screening strategies. We have utilized the CENP-E&#160;knockout cell line to establish a novel approach to validate the specificity and toxicity of the CENP-E inhibitors, which is useful for drug development and biological research.

The CRISPR (clustered regularly interspaced short palindromic repeats)/Cas9 system has emerged as a powerful tool for precise and efficient gene editing ...