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

Summary

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.

Abstract

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.

Introduction

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 DNA^6,7,8 and has become one of the most widely used tools for gene-editing to date^9,10,11,12. 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 aegypti^13,14, 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/Cas9^17,18,19. 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 references^17,18,19.

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.

Protocol

1. Target design and in vitro synthesis of sgRNA

1. 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.
2. 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.
3. 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' 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).
4. Use the commercially available gRNA synthesis kit to generate the designed sgRNA. Perform each step following the user's guide. Resuspend the gRNA product in nuclease-free water, quantify the concentration using a UV-Vis spectrophotometer, and store at -80 °C prior to use (the concentration of successfully synthesized single gRNA [10 µL] with the kit used here is about 4000-6000 ng/µL, or even higher).
   ​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.

2. Embryo collection and preparation

1. Place B. dorsalis pupae 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 °C, and a 14:10 L/D cycle(lights on at six in the morning, lights off at eight in the evening).
2. 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 (<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.
3. 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' survival rate.
4. 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 <10% RH for 10 min is optional. Dip the embryos into halocarbon oil during injection to avoid further desiccation (Figure 2C).

3. Microinjection of the embryo

1. 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.
2. Prepare the working solution. Mix the Cas9 protein and the corresponding sgRNA to the following working concentrations: sgRNA, 300 ng/µL; Cas9 protein, 150 ng/µL. Add 1 µL 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/µL; 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 editing^17,19. 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.
3. 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.
4. Add 3 µL of the mixture into the injection needle. Avoid introducing air bubbles that can possibly clog the needle. Open the needle using a Microgrinder.
5. Connect the needle to the micromanipulator according to the user guide (Figure 2E).
6. 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.
7. 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.
   ​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.

4. Post-injection insect rearing

1. Put the injection plate with the injected embryos into an artificial climate chamber kept at 55% RH, 26.5 °C, and a 14:10 L/D cycle(lights on at six in the morning, lights off at eight in the evening).
2. 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.
   ​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 (<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.
3. 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.

5. Mutant screening

1. 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.
2. 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.
3. 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.
4. Perform PCR using the following cycling conditions: 98 °C for 3 min, followed by 35 cycles of 98 °C for 10 s, 15 s at an annealing temperature appropriate for the designed primers, 72 °C for 35 s, followed by a final extension of 72 °C for 10 min.
5. 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.

Results

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...

Discussion

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...

Materials

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