Presented here is a protocol of Helicoverpa armigera (Hübner) 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 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.
The application of genome editing technology provides an efficient tool to achieve target-gene mutants in diverse species. The emergence of the clustered regularly interspaced short palindromic repeats (CRISPR)/associated protein 9 (Cas9) system provides a novel method to manipulate genomes1. The CRISPR/Cas9 system consists of a guide RNA (gRNA) and the Cas9 endonuclease2,3, while the gRNA can be further divided into two parts, a target complementary CRISPR RNA (crRNA) and a trans-activating crRNA (tracrRNA). The gRNA integrates with Cas9 endonuclease and forms a ribonucleoprotein (RNP). With the gRNA, Cas9 endonuclease can be directed to a specific site of the genome via base complementation. The RuvC and HNH domains of the Cas9 cleave the target site of the genome three bases before the protospacer-adjacent motif (PAM) sequence and create a double-strand break (DSB). The DNA cleavage can then be repaired through two mechanisms, non-homologous end joining (NHEJ) or homology-directed repair (HDR)4. Repair of the DSB introduces insertions or deletions as a way to inactivate the targeted gene, potentially causing a complete loss of gene function. Hence, the hereditable and specificity of the CRISPR/Cas9 system make it a robust method to characterize gene functions in vivo and analyze gene interactions5.
With numerous merits, the CRISPR/Cas9 system has been applied to various fields, including biomedicine6,7, gene therapy8,9, and agriculture10,11,12, and has been used for various biological systems including microorganisms13, plants14,15, nematodes16 , and mammals17. In invertebrates, many insect species have been subjected to CRISPR/Cas9 genome editing, such as the fruit fly Drosophila melanogaster and beyond18,19,20,21,22.
Helicoverpa armigera is one of the most destructive pests worldwide23, and damages numerous crops, including cotton, soybean, and sorghum24,25. With the development of sequencing technology, the genome of H. armigera, as well as that of a range of Lepidoptera insect species, have been sequenced completely26,27,28,29. A large number of resistance and olfactory receptor genes have been identified and characterized from these insects in recent years19,27,28,29. Some resistance-related genes have been identified in H. armigera, such as the genes encoding for cadherin30, an ATP-binding cassette transporter31,32, as well as HaTSPAN133. Knockout of these genes using CRISPR/Cas9 technology results in a high level of resistance to Bacillus thuringiensis (BT) toxin in susceptible strains. Also, Chang et al. (2017) knocked out a pheromone receptor, which validated its significant function in mating time regulation19. These reports suggest that CRISPR/Cas9 can act as an effective tool to study gene function in vivo in insect systems. However, a detailed procedure for CRISPR/Cas9 modification in insect systems remains incomplete, which limits its application range in insect functional genomics.
Here, we present a protocol for knocking out a functional gene in H. armigera using the CRISPR/Cas9 system. A detailed step-by-step protocol is provided, including the design and preparation of gene-specific primers for gRNA production, embryo collection, microinjection, insect rearing, and mutant identification. This protocol serves as a valuable reference to manipulate any functional genes in H. armigera and can be extended to other Lepidoptera species.
1. Design of gene-specific primers and preparation of sgRNA
2. Embryo preparation and collection
3. Microinjection of embryos
4. Post-injection insect rearing
5. Knock-out mutant detection
This protocol provides detailed steps for obtaining gene knock-out lines of H. armigera using CRISPR/Cas9 technology. The representative results obtained by this protocol are summarized for gDNA selection, embryo collection and injection, insect rearing, and mutant detection.
In this study, the target site of our gene of interest was located in its second exon (Figure 2A). This site was highly conserved, and the target band fragment of synthesized sgRNA was confirmed using agarose gel electrophoresis (Figure 2B,C,D).
The male and female moths were initially reared in separate net cages to prevent mating ahead of schedule and to ensure a sufficient quantity of embryos as much as possible. In general, a total number of 300 fertilized eggs were collected and were immediately injected with the sgRNA/Cas9 protein mixture (300-500 ng/µL of sgRNA, 200 ng/µL of Cas9 protein) at the one-cell stage. The injection volume was about one-tenth that of the embryos. After microinjection, the embryos were reared as described in section 4, and 40%-60% of injected embryos survived.
The mutant detection of a single sgRNA target was performed by sequencing the PCR products from G1 parental adults (Figure 6B). We also tested the effectiveness of using non-overlapping sgRNA pairs across different exons. The large deletion of the mutants (Figure 6C,D) can be easily distinguished from wild type bands (Figure 6A).
The mutation rate calculated in this protocol was 87.50% when 16 individuals are randomly tested, indicating that this protocol was highly-efficient. Gene knockout results were shown in several genotypes, but the majority of mutants identified from our screening were -2 bp type. Mutations resulted in the premature termination of protein translation in the genome, which subsequently led to the loss of gene function.
Figure 1: The flowchart for the preparation of sgRNA. Please click here to view a larger version of this figure.
Figure 2: Selection and synthesis of target sgRNAs from H. armigera. (A) The yellow domain represents the exon, while the black line represents the intron. The red sequences indicate the target sequence, and the blue sequences indicate the protospacer adjacent motif (PAM). (B) PCR assembly of the sgRNA DNA template. (C) The in vitro transcription product. (D) Purification of sgRNA. Please click here to view a larger version of this figure.
Figure 3: Embryo collection. (A) A net cage covered with black cloth. The male and female moths of H. armigera were mating. (B) The microscope slide without embryos. (C) The microscope slide containing 50-100 embryos on pieces of black cloth. Please click here to view a larger version of this figure.
Figure 4: Needle preparation. (A) Micropipette puller. (B) Tip of a microinjection needle after pulling by a micropipette puller. The dotted box indicates the magnified needle tip. Scale bar represents 1 mm. Please click here to view a larger version of this figure.
Figure 5: Embryo microinjections. (A) The whole set of a microinjection system containing a microscope (middle) and an electronic microinjector (left) connected to a micromanipulator (right). (B) Embryos and microinjection needle. (C) The injection site of the embryo is labeled with the red arrow. Scale bar represents 200 µm. (D) A hatched larva under the microscope. Scale bar represents 1 mm. Please click here to view a larger version of this figure.
Figure 6: Detection of mutants by PCR and gel electrophoresis. The black arrows and red lines indicate the target sites of the sgRNA. (A) The band in lane 1 represents the amplification fragment derived from wild type. (B) The bands in lane 2 and 3 represent the amplification fragment derived from mutant using a single sgRNA target. (C) The detection of a heterozygote using a pair of non-overlapping sgRNA. The bands in lanes 4 and 5 represent the amplification fragment derived from the mutation of two sgRNA targets. The lower bands indicate a large fragment deletion. (D) The results are derived from a homozygote. The bands in lane 6 and 7 indicate the large fragment deletion. Please click here to view a larger version of this figure.
The application of the CRISPR/Cas9 system has provided powerful technical support for the analysis of gene function and interaction among various genes. The detailed protocol we present here demonstrates the generation of a homozygote mutant in H. armigera via CRISPR/Cas9 genome editing. This reliable procedure provides a straightforward way for directed gene mutagenesis in H. armigera.
The choice of CRISPR target sites could affect the mutagenesis efficiency37. In this protocol, we compared and analyzed multiple results from the online website CRISPOR to obtain an appropriate target site. In silico, gRNA predictions present some advantages. Firstly, they analyze the whole genome of H. armigera when designing sgRNAs to minimize the off-target effects. The online resources mentioned above, as well as CHOPCHOP (http://chopchop.cbu.uib.no/), function with a number of Lepidoptera genomes, which could be beneficial for gene editing in other Lepidopteran moths. Secondly, the ranking of the candidate sgRNAs directly compares possibilities but might include some variations based on the different algorithms. The candidate sequence with high ratings in both lists tends to be more reliable. However, a major limitation of this protocol is that a large number of insect genomes are absent in the databases of the websites, so there is potential for off-target effects. Another limitation is that the PAM sequence is necessary for the sgRNA design, which may result in the inability to find an appropriate target site.
The tissues used for mutant screening are also a crucial factor. The survival rate, life cycle, and physiological functions of insects should not be affected. In our process of exploring the optimal gDNA extraction method, the micro-hemolymph extraction from larvae was attempted for mutant detection to save time and avoid mating (unpublished data). However, this method brought more challenges regarding the efficiency of PCR amplification and the survival rates of adult (data not shown). In addition, Zheng et al.38 reported a non-destructive method for gDNA extraction using the exuviate or puparia. Based on those findings, we modified and explored an approach using hind legs for gDNA extraction, which allows adult moths to survive and mate naturally, significantly improving the detection accuracy of a given genotype. Therefore, we developed a new method to increase the success rate of gDNA extraction by removing one of the hind legs from each adult candidate. We further confirmed that this operation did not affect the survival rate and mating frequency of adult moths. Furthermore, we found that the large fragment deletion can be easily observed by the gel electrophoresis when co-injected with a pair of gRNAs across exon-regions (Figure 6), which simplifies the process of mutant identification when screening.
The eggs of H. armigera are collected on a black cloth, which makes it easy to distinguish the eggs under the microscope in the process of microinjection (Figure 5B). Due to the common reproductive behaviors of Lepidopteran moths such as mating, oviposition, hatching, and eclosion39,40,41, this egg-collecting technique could also be applied for other Lepidopteran moths.
In conclusion, the CRISPR/Cas9 system has proven to be a reliable tool for facilitating functional genomics studies in H. armigera. The step-by-step descriptions enable users to complete an integral gene-editing process.
The authors have nothing to disclose.
This work was supported by National Natural Science Foundation of China (31725023, 31861133019 to GW, and 31171912 to CY).
2kb DNA ladder | TransGen Biotech | BM101 | |
Capillary Glass | World Precision Instrucments | 504949 | referred to as "capillary glass" in the protocol |
Double Sided Tape | Minnesota Mining and Manufacturing Corporation | 665 | |
Eppendorf FemtoJet 4i Microinjector | Eppendorf Corporate | E5252000021 | |
Eppendorf InjectMan 4 micromanipulator | Eppendorf Corporate | 5192000051 | |
Eppendorf Microloader Pipette Tips | Eppendorf Corporate | G2835241 | |
GeneArt Precision gRNA Synthesis Kit | Thermo Fisher Scientific | A29377 | |
Microscope Slide | Sail Brand | 7105 | |
Olympus Microscope | Olympus Corporation | SZX16 | |
PrimeSTAR HS (Premix) | Takara Biomedical Technology | R040 | used for mutant detection |
Sutter Micropipette Puller | Sutter Instrument Company | P-1000 | |
TIANamp Genomic DNA Kit | TIANGEN Corporate | DP304-03 | |
TrueCut Cas9 Protein v2 | Thermo Fisher Scientific | A36499 |