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Review

CRISPR/Cas9 Genome Editing in the Diamondback Moth: Current Progress, Challenges, and Prospects

by
Muhammad Asad
1,2,3,4,5,†,
Yanpeng Chang
1,2,3,4,5,†,
Jianying Liao
1,2,3,4,5 and
Guang Yang
1,2,3,4,5,*
1
State Key Laboratory of Agricultural and Forestry Biosecurity, Institute of Applied Ecology, Fujian Agriculture and Forestry University, Fuzhou 350002, China
2
Joint International Research Laboratory of Ecological Pest Control, Ministry of Education, Fuzhou 350002, China
3
Ministerial and Provincial Joint Innovation Centre for Safety Production of Cross-Strait Crops, Fujian Agriculture and Forestry University, Fuzhou 350002, China
4
Key Laboratory of Integrated Pest Management for Fujian-Taiwan Crops, Ministry of Agriculture, Fuzhou 350002, China
5
Key Laboratory of Green Pest Control, Fujian Province University, Fuzhou 350002, China
*
Author to whom correspondence should be addressed.
†
These authors contributed equally to this work.
Int. J. Mol. Sci. 2025, 26(4), 1515; https://doi.org/10.3390/ijms26041515
Submission received: 2 January 2025 / Revised: 4 February 2025 / Accepted: 10 February 2025 / Published: 11 February 2025
(This article belongs to the Special Issue Recent Progress in Molecular Biology of RNA 2.0)
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Abstract

:
The development of site-specific genome-editing tools like CRISPR (clustered regularly interspaced short palindromic repeat) and its associated protein, Cas9, is revolutionizing genetic engineering with its highly efficient mechanism, offering the potential for effective pest management. Recently, CRISPR/Cas9 gene-editing has been extensively utilized in the management of the diamondback moth, Plutella xylostella (L.), a highly destructive pest of vegetable crops, for different purposes, such as gene function analysis and genetic control. However, the progress related to this gene-editing tool in P. xylostella has not yet been summarized. This review highlights the progress and applications of CRISPR/Cas9 in uncovering the genes critical for development, reproduction, and insecticide resistance in P. xylostella. Moreover, the progress related to the CRISPR/Cas9 gene drive for population suppression and modifications has also been discussed. In addition to the significant progress made, challenges such as low germline editing efficiency and limited homology-directed repair remain obstacles to its widespread application. To address these limitations, we have discussed the different strategies that are anticipated to improve the efficiency of CRISPR/Cas9, paving the way to it becoming a pivotal tool in sustainable pest management. Therefore, the present review will help researchers in the future enhance the efficiency of the CRISPR/Cas9 system and use it to manage the diamondback moth.

1. Introduction

The diamondback moth (DBM), Plutella xylostella (L.) (Lepidoptera: Plutellidae), is a widespread pest species within the Lepidopteran group that causes significant damage to cruciferous crops worldwide [1]. P. xylostella is known as an oligophagous pest as it feeds on both wild and cultivated cruciferous vegetables (Cruciferae), including some commercially and economically significant vegetables like cauliflower, cabbage, and canola [2]. The damage caused by P. xylostella, along with management costs, results in estimated global economic losses of USD 4–5 billion annually [1]. The life cycle of the P. xylostella varies with temperature, ranging from 4 to 20 generations per year [1,2,3]. First-instar larvae act as leaf miners, feeding on the spongy tissue of leaves. As they progress to later instars (second, third, and fourth), the larvae transition to feeding on the leaf surface, consuming the outer green layer, buds, flowers, stems, and seeds (Figure 1). This feeding behavior leads to significant crop damage, reducing both yield and quality of crops [1,2]. Insecticides have traditionally been the main method for controlling P. xylostella populations. However, the moth has evolved resistance to almost all major groups of insecticides, making it challenging to manage in agricultural settings [1,4,5]. Given these challenges, there is a critical need for innovative and effective strategies, such as genetic-based control methods [6]. These include RNA interference (RNAi) for gene silencing, transgenic approaches like crops expressing Bacillus thuringiensis (Bt) toxins, and clustered regularly interspaced short palindromic repeat and its associated protein Cas9 (CRISPR/Cas9) for precise genome editing, all of which offer sustainable alternatives to chemical pesticides [6]. Among these strategies, the CRISPR/Cas9 system is known to be a promising tool to develop advanced pest control strategies, including gene drive systems for population suppression [7,8] and genetic sterile insect techniques (gSIT) such as the precision-guided sterile insect technique (pgSIT) [9,10].
As early as 1987, the structure of CRISPR was first identified in bacterial species known as Escherichia coli [11]. However, the same CRISPR structures were further discovered in other various bacterial species, and the abbreviation of this system was developed in 2002 [12]. CRISPR gene loci are mainly located adjacent to Cas9 genes and typically comprise multiple noncontiguous direct repeats that have been divided by flexible sequences referred to spacers [13]. Since 2013, the CRISPR/Cas system has been commonly employed in different types of biological research, and CRISPR/Cas9 is the most utilized variant of this system [14,15]. The CRISPR/Cas9 gene-editing system has dramatically improved the field of genetic engineering due to its accurate and highly efficient mechanism [16]. The guide RNA (gRNA) is an artificial synthesized RNA molecule that is complementary to the target sequence in the genome of an organism and is essential for the proper functioning of this system. Cas9 proteins are known as molecular scissors, which are paired with the gRNA. The Cas9–gRNA complex initiates the precise double-stranded break (DSB) in the target DNA sequence after locating it in the genome (Figure 2). After the DSB, the biological mechanisms of DNA repairing initiate, including non-homologous end joining (NHEJ) and homology-directed repair (HDR) [17,18,19]. NHEJ frequently triggers the minor deletions or insertions known as indels, which eventually disrupt the open-reading frame and result in gene knockout [18]. On the contrary, HDR can be used to create targeted genetic alterations by introducing a DNA template alongside the CRISPR/Cas9 components, facilitating accurate gene editing with the desired alterations [18,20,21,22]. Due to its efficiency and specificity, the CRISPR/Cas9 system has been effectively used in various organisms, including plants [23], microbes [24], animals [25], insects [26,27], and humans [28].
Moreover, this system presents an optimistic method for pest control within the field of pest management. This technique involves targeting certain genes that are crucial for the survival or reproduction of pests and that may be used to develop transgenic crops, which are resistant to pest attack, by altering the genes associated with susceptibility to pests, thereby decreasing the need for chemical pesticides and encouraging sustainable agriculture practices. In addition, the use of the CRISPR/Cas9 system has been employed to investigate the drug resistance mechanisms in pathogens, demonstrating its promise in tackling issues linked to pest and pathogen control. CRISPR/Cas9 technology shows several benefits, such as its adaptability and effectiveness in facilitating gene editing in different organisms [20]. Moreover, it has been demonstrated that ribonucleoprotein complexes and DNA-free genome editing reduce off-target effects, ultimately improving the accuracy of gene editing through this technology [29]. In comparison to prior gene-editing technologies, CRISPR/Cas9 exhibits a notable degree of efficiency and cost-effectiveness, thereby expediting advancements in the fields of research and development.

2. Applications of CRISPR/Cas9-Based Gene Knockout System in Diamondback Moth

The emergence of CRISPR/Cas9-based gene editing has shown great potential in pest management applications. Before its emergence, research on the gene function in the P. xylostella was limited due to the inefficacy of RNAi and the absence of direct genome-editing technology. In 2016, the effective use of CRISPR/Cas9 to specifically target Abdominal A gene (Pxabd-A) in P. xylostella validated the promise of utilizing this approach to disturb the genes in agricultural pests [30]. Subsequently, by using this system in P. xylostella, the understanding of genes associated with growth, development, and reproduction gradually improved, leading to significant advancements in pesticide and Bt resistance mechanisms (Table 1).

2.1. Development and Reproduction

CRISPR/Cas9 technology has significant potential as a tool for gene function study, which has effectively been used to uncover the roles of genes related to development and reproduction in the diamondback moth. Vitellogenin receptor (VGR) and Vitellogenin (Vg) genes play crucial functions in insect oocyte maturation and embryonic development. Disruption of PxVg and PxVgR through the CRISPR/Cas9 system leads to a reduction in ovary size and decreased hatching rates, while egg production remains unaffected [32,33]. Serine protease 2 (Ser2) belongs to the Serine proteases gene family and serves as a crucial constituent of insect seminal fluid proteins. A mutation in PxSer2 leads to a dominant form of male infertility that is inherited by the next generation [34]. Vitelline membrane proteins (VMPs) are key components involved in the formation of the insect vitelline membrane layer, which is essential for oogenesis and embryonic development. Knockout of PxVMP26 leads to abnormal eggs with a lower hatching rate, while egg production remains unaffected [35]. Piwi protein plays a pivotal role in animal germ cell development and transposon silencing through its interaction with piRNA-associated small non-coding RNAs. Knockout of PxPiwi gene prolongs the pupal stage, leading to abnormal eclosion [36]. Deletion of biogenic amine tyramine receptor 1 (PxTAR1) inhibits sex pheromone biosynthesis and ovarian development, resulting in reduced mature oocytes and decreased egg production [37]. Ovary-serine protease (Osp) is crucial for the process of oogenesis and the development of ovaries. The absence of PxOsp results in recessive infertility in females, whereas male fertility stays unaffected [38].
Utilizing CRISPR/Cas9 in the P. xylostella not only reveals the conservation of genes typically involved in reproductive development but also sheds light on the participation of non-traditional reproductive development genes. For example, the transcription factors (TFs) may have significant functions in growth and development, sexual dimorphism, and pesticide resistance. Disruption of the Homeobox transcription factor gene cluster leads to a reduction in the egg hatching rate [31]. Dicer, a member of the RNase III family endonucleases, shows precise recognition and cleavage of dsRNA. Knockout mutants of PxDcr-1 display higher mortality rates, lower eclosion rates, reduced egg production, decreased hatching rates, and ovarian abnormalities, whereas no significant differences are seen between PxDcr-2 knockout mutants and wild-type individuals in terms of their developmental and reproductive characteristics [39].

2.2. Pigmentation

The study of phenotypic marker genes is essential to screening transgenic insects for the development of specific genetic control systems. When modeling gene drives and assessing their effectiveness on target insects, it is crucial to initially target endogenous phenotypic genes. Knockout of PxYellow causes modified body pigmentation across larvae, pupae, and adults but does not cause significant differences in adult oviposition or egg hatching rate. This suggests that PxYellow does not have direct role in growth, development, and reproductive regulation, indicating its potential for use as a genetic marker for development of genetic control technologies [42]. The eye color gene has been served as an obvious marker in many prototype strategies for genetic control of insects. CRISPR/Cas9-based disruption of kynurenine 3-hydroxylase and Cardinal in the diamondback moth results to yellow compound eyes in adults, along with changes in brain, ovary, and testis pigmentation [40]. Ebony gene serves as a core enzyme involved in dopamine conversion to N-β-alanyldopamine and plays an important function in the melanin production. Disruption of PxEbony results in the development of dark pigmentation in the bodies of larvae, pupae, and adult insects. Additionally, there is a notable decrease in the rates of egg hatching and the survival of larvae when compared to the wild-type strain [41].

2.3. Sex Determination

Insects have developed intricate and complex mechanisms for sex determination, showing significant variations. Presently, the procedures for determining sex have been thoroughly studied in Dipteran insects such as Drosophila. Nevertheless, there is a lack of research on the processes of sex determination in Lepidopteran insects. However, the use of the CRISPR/Cas9 system has allowed for the study of the partial sex determination mechanism in the lepidopteran model insect B. mori. For P. xylostella, only a few genes related to sex determination pathways have been studies by utilizing the CRISPR/Cas9 genome-editing tool. Doublesex (Dsx) is a pivotal gene in insects that is essential for sex determination. It plays a critical part in the process of sexual differentiation and development. In the diamondback moth, the Dsx gene possesses a single transcript for males and three transcripts for females. The CRISPR/Cas9 method has been utilized to deliberately introduce mutations in both the male- and female-specific isoforms, as well as independently targeting each particular isoform for males and females. The male mutants exhibits the expression of female-specific transcripts, while the female mutants showed the expression of male-specific transcripts, ultimately leading to gender-specific sterility [43]. The P-element somatic inhibitor (PSI) is crucial for promoting male-specific splicing of the Dsx gene, which is essential for male sexual differentiation. Knockout of PxPSI results in the malformation of male reproductive organs, leading to infertility [44]. Seminal fluid proteins (SFPs) are major determinants of reproductive success between sexes. Knockout of one SFPs gene(PxSast1) reduces male mating ability and fertilization capacity by suppressing the expression of female follicular genes, leading to defects in oogenesis [45].

2.4. Circadian Rhythms

The biological clock regulates the rhythmicity of metabolism, physiology, and behavior in organisms. The majority of organisms possess a circadian rhythm that is regulated by a molecular oscillator consisting of multiple clock genes that are constant across species. These clock genes include timeless (tim), period (per), cryptochrome (cry), clock (clk), and cycle (cyc) [71,72]. The absence of PxCry1 disrupts rhythmic locomotion under light–dark conditions and eliminates rhythmicity under continuous darkness. This highlights that PxCry1 is crucial for the regulation of diurnal rhythm in the diamondback moth [46]. Deletion of PxCry2 or Pxper eliminates the peak activity after the absence of light during light–dark cycles. Furthermore, the lack of PxCry2 diminishes overall activity, which is essential for sustaining internal rhythms in a state of continuous darkness [48]. Phototaxis, the movement in response to light, is an important ecological trait for nocturnal insects. Three opsin genes, namely, LW-opsin, BL-opsin, and UV-opsin, have been identified in the genome of the diamondback moth. These genes are mainly expressed in the head region, with LW-opsin having the highest expression level. Knockout mutants lacking PxLW-opsin show reduced phototaxis towards various white light wavelengths and impaired perception of white light, green light, and infrared light [47].

2.5. Ecological Adaptability

Insects play a critical role in ecosystems by occupying essential ecological niches, largely due to their impressive capacity for adaptation. This adaptability is evident in their ability to thrive in various environments through host adaptation, which involves adjustments in diet, reproductive strategies, and behavior. A key impetus for the evolutionary adaptation of herbivorous insects is the necessity to overcome the defense mechanisms of host plants, thereby optimizing their ability to acquire necessary nutrients for survival and reproduction. Glucosinolate sulfatases (GSS) are recognized as pivotal adaptive mechanisms employed by P. xylostella to counteract the glucosinolate–myrosinase defense systems present in cruciferous plants [73,74]. PxGSS1 functions by secreting a desulfurization inhibitor that hinders the formation of harmful compounds derived from the standard plant defensive molecule 4-(methylsulfinyl) butyl glucosinolate (4MSOB-GL). A knockout of PxGSS1 leads to a reduced capacity for host adaptation, elevated larval mortality rates, and prolonged pupal stage. Conversely, mutation in PxGSS2 reduces egg hatching rate, increases larval mortality, and prolongs pupal development. Notably, a loss-of-function mutation in PxGSS3 does not exert a significant effect on host adaptability in P. xylostella [49,50,52,75]. Similarly, knockout of the methyltransferase-like 14 gene (Pxmettl14), which encodes an RNA transferase gene, markedly impairs host adaptability. This impairment is evidenced by significantly reduced pupal weight, oviposition rate, and egg hatching rate in comparison to the wild type when transitioning from artificial feed to host plants [53]. The upregulation of the glycoside hydrolase family 1 (GH1) member Px008848 protein reduces the survival of larvae when they feed on host plants. Conversely, the knockout of Px008848 greatly increases the survival rate of mutant larvae relative to wildtype [50]. Isothiocyanates (ITCs) exhibit both repellent and toxic effects on herbivores. Specialist herbivores, however, utilize volatile ITCs as essential cues for locating the host. The olfactory receptors Or35 and Or49 play a significant role in this process. CRISPR/Cas9-mediated mutations in either PxOr35 or PxOr49 lead to a decrease in oviposition preference for ITCs, whereas double knockout strains of female adults completely lose their preference for ITCs and show no selective preference between wild-type Arabidopsis and ITC-knockout Arabidopsis plants [51]. Furthermore, PxOr16 demonstrates specific sensitivity to heptanal, an epidermal volatile compound released by Cotesia vestalis. Knockout of PxOr16 eliminates the avoidance behavior towards heptanal [56].
Climate change presents novel challenges for temperature-adopting organisms, particularly insects, in relation to global warming. Trehalose acts as the main source of sugar in the hemolymph of insects and has a crucial function in energy metabolism and stress response. In the diamondback moth, knockout of the PxTret1-like gene significantly reduces tolerance to extreme temperatures, decreases reproductive capacity, enhances the mortality rate, and hinders development, reproduction, and temperature adaptability [55]. Very-long-chain fatty acids (VLCFAs) are essential for insects to adapt to thermal stress. The 3-hydroxy acyl-CoA dehydratase 2 gene (Hacd2) is known as a crucial enzyme in VLCFA synthesis. Knockout of PxHacd2 results in increased cuticular permeability caused by decreased VLCFA levels in P. xylostella. This, in turn, makes them less capable of adapting to dry environmental conditions, leading to significantly reduced survival rates and reproductive capacity [54]. Knockout of peripheral genes Karyopherin subunit beta 1 (KPNB1) and Toiled-coil domain containing 2 (GCC2) weakens the response ability of P. xylostella with respect to extreme temperatures while concurrently inhibiting the development and reproduction processes [57].

2.6. Insecticide Resistance

The issue of insecticide resistance in P. xylostella has significantly escalated, with almost all chemical insecticides showing various levels of resistance. The lack of understanding about the mechanisms underlying this resistance has impeded the advancement of basic strategies to tackle this issue. However, by using CRISPR/Cas9 technology on the P. xylostella, targeted mutations have enabled the comprehensive investigation of the mechanisms responsible for its insecticide resistance. Research on the gene mutations associated with resistance has established a foundation for the further elucidation of how pests develop resistance against chemical insecticides. The main target of diamide insecticides is the ryanodine receptor (RyR). Previous studies have identified three specific mutations at positions—i.e., G4946E, I4790K, and I4790M—in RyR that are associated with diamide resistance. Two knock-in strains, PxI4790K-KI and PxI4790M-KI, have been developed using a CRISPR/Cas9-mediated system, with a single-stranded oligonucleotide DNA (ssODNA) of approximately 120 base pairs serving as the knock-in template. Compared to susceptible strains, PxI4790K-KI exhibits significantly increased resistance to cyantraniliprole and chlorantraniliprole, as well as to the diamide flubendiamide. Similarly, PxI4790M-KI demonstrates enhanced resistance to chlorantraniliprole, cyantraniliprole, and flubendiamide, [64,67]. Various kinds of insecticides have the nicotinic acetylcholine receptor (nAChR) as a common target. A CRISPR/Cas9-mediated deletion of PxnAChRα6 confers significantly increased resistance to Spinosad and spinetoram. However, this mutation does not result in notable changes; it causes only limited increases in resistance to abamectin, imidacloprid, indoxacarb, β-cypermethrin, chlorantraniliprole, and metaflumizone [63]. Cyclopentadiene organochlorine and phenylpyrazole insecticides inhibit neurotransmission in insects by blocking the GABA-gated chloride channel. and it has been confirmed that the A301S mutation in GABA receptors contributes to cyclodiene, organochlorine, and phenylpyrazole pesticide resistance. CRISPR/Cas9-mediated editing of PxGABARalpha in an insecticide-susceptible strain of P. xylostella does not confer resistance to dieldrin, endosulfan, or fipronil [58]. Knockout studies of two fipronil-sensitive and fipronil-resistant genes (PxRdl1 and PxRdl2) indicate that disruption of PxRdl1 diminishes the effectiveness of fipronil, while disrupting the fipronil-resistant PxRdl2 increases the effectiveness of fipronil [76]. The glutamate-gated chloride channel (GluCl) is a primary target for avermectin insecticides. A mutation from valine to isoleucine (V263I) in GluCl has been confirmed to involve varying degrees of resistance for abamectin in the diamondback moth. A CRISPR/Cas9-edited strain containing the homozygous V263I mutation (PxGluClV263I-KI) has been developed, which exhibits significant resistance to avermectin. However, this mutation also results in a notable reduction in reproductive capacity [66].
The fast development of insecticide resistance in pests poses a substantial obstacle to the sustainable utilization of biological pesticides and genetically engineered crops that generate Bacillus thuringiensis (Bt) toxins. The development of a strong resistance to Bt toxins, specifically Cry1Ac, in lepidopteran insects like the diamondback moth has been associated with mutations or the reduced activity of ABC transporter subfamily C genes (ABCC2 and ABCC3). CRISPR/Cas9-mediated knockout strain (PxABCC2-KO) demonstrates a 724-fold increase in resistance, whereas in contrast to the susceptible strain, the PxABCC3-KO strain shows a 413-fold higher resistance to the Cry1Ac protoxin. Both knockout strains exhibit a substantial decrease in the binding affinity between the Cry1Ac toxin and midgut membrane vesicles compared to the sensitive strain. Furthermore, the loss of these two genes leads to a resistance that is more than 8000 times higher than Cry1Ac and 380 times higher than Cry1Fa. This highlights the fact that ABC transporters serve as redundant toxin receptors in the diamondback moth [51,59,62]. The use of CRISPR/Cas9 technology to simultaneously knockout the PxAPN1 and PxAPN3a results in a remarkable 1425-fold increase in resistance to the Cry1Ac toxin. Furthermore, the simultaneous knockout of four genes, namely, PxABCC2, PxABCC3, PxAPN1, and PxAPN3, results in an extraordinary increase of over 34000-fold in resistance against Cry1Ac toxin [60]. The polycalin protein is known as a possible receptor for Bt toxin, and knockout of the PxPolycalin gene leads to a decline in the resistance of the diamondback moth to Cry1Ac toxin [61]. Methionine aminopeptidase (MetAP) plays a crucial role in peptide synthesis, and knockout of PxMetAP contributes to increased resistance against Cry1Ac toxin in P. xylostella [65]. Juvenile hormone binding protein (JHBP) is a pivotal regulatory factor in the signaling pathway of juvenile hormone (JH). Notably, a mutation within PxJHBP leads to a substantial increase in sensitivity to Cry1Ac, resulting in a shortened lifespan and significantly impaired reproductive capacity [68].

3. Application of CRISPR/Cas9 Knock-In and Gene Drive Systems in the Diamondback Moth

Gene drive technology holds the potential to rapidly disseminate desirable traits into wild populations, presenting an efficient method for controlling or eradicating target species [77,78,79]. With the continuous advancement of gene-editing techniques, gene drive systems have become a promising approach for pest control applications. Among these approaches, CRISPR/Cas9-based gene drive systems have gained considerable interest because of their simplicity, versatility, and high efficacy [8,80,81]. The successful implementation of CRISPR/Cas9-based gene drives in dipteran insects, specifically in fruit flies and mosquitoes, has opened the possibility of developing a functional gene drive in the diamondback moth. Two types of CRISPR/Cas9-based gene drive systems, namely, the global gene drive system and the split-drive system, have been constructed to modify the population of diamondback moth (Figure 3). In the global gene drive system, all gene drive components, including the Cas9 gene, gRNA, and marker gene, are integrated into a single construct (Figure 3). Specifically, a global gene drive system in P. xylostella has been successfully developed, targeting PxYellow, known as a phenotypic marker gene, and gene drive components are precisely integrated into the diamondback moth genome, which are inherited by subsequent generations. The HDR-based efficiency of the gene drive ranges from 6.67% to 12.59%, demonstrating the possibility of developing effective CRISPR/Cas9-based pest control strategies for the diamondback moth. Additionally, it provides a solid foundation for constructing a functional CRISPR/Cas9 gene drive for diamondback moth [69]. In split-drive system, the Cas9 gene along with one marker gene is inserted into autonomous chromosomes using the piggyBac-mediated transposition system, while the gRNA and another marker (drive component) are inserted into the target gene through the CRISPR/Cas9 knock-in system. Two distinct lines, one producing Cas9 and the other producing gRNA, are created and crossed to ensure the inheritance of the drive components in subsequent generations (Figure 3). The Cas9 components are inherited normally, whereas the gRNA components bypass Mendelian segregation. A split-drive gene drive system targeting two phenotypic genes, PxKmo and PxYellow, has been developed in P. xylostella. In this system, different endogenous promoters are tested to drive Cas9 expression, alongside a single gRNA, which results in a high level of cleavage in somatic cells but low cleavage in germline cells and no homing effects. This study lays the groundwork for improving the functional split-drive system, which can be further utilized to develop genetic control methods for P. xylostella [70]. In addition to gene drive systems, the CRISPR/Cas9 knock-in method has been utilized to evaluate its ability to cleave DNA in the diamondback moth cell lines and to enable the targeted insertion of marker genes into sex chromosomes for the purpose of separating genders at an early stage. A transgenic diamondback moth cell line expressing Cas9 has been successfully generated using two ubiquitous promoters: Px217-2 and IE1. The Px217-2 promoter exhibits robust gene knockout efficiency compare to the IE1 promoter in diamondback moth cell lines [82]. The use of CRISPR/Cas9 technology to precisely insert marker genes into sex chromosomes allows for the separation of genders, providing a secure and economically efficient approach that may be applied to the mass production of transgenic females or males. Using CRISPR/Cas9 knock-in technology, the DsRed2 gene has been inserted into the W chromosome at a specific genomic locus. However, this insertion severely affects the typical growth and reproduction of the diamondback moth, leading to deformities and infertility, indicating that this particular genomic locus is unsuitable for integrating exogenous sequences. Conversely, the cyan fluorescent protein (CFP), along with the Cas9 gene (Cas9-CFP), has been precisely integrated into another genomic locus on the Z chromosome. The introduction of the Cas9 gene along with CFP marker gene into the Z chromosome enables gender separation through fluorescence labeling. These findings demonstrate the feasibility of integrating exogenous genes into different genomic sites in the diamondback moth [83].

4. Challenges and Possible Solutions of CRISPR/Cas9 Gene-Editing in the Diamondback Moth

Although CRISPR/Cas9 has made significant progress in the genome editing of the diamondback moth, it is crucial to acknowledge and address the limitations associated with this technology. These limitations include the possibility of off-target mutation, the delivery of CRISPR/Cas9 components, variations in knockout efficiencies, low germline cleavage, and limited homology-directed repair, which need to be addressed in order to further improve this technology.
The potential for off-target mutations arises from mismatches between sgRNA and its target site, resulting in unintended and nonspecific changes in the genome. The effectiveness of CRISPR/Cas9 depends on the 20 base pair nucleotide sequences of the gRNA and the PAM sequences located close to the target site [19]. Studies have demonstrated that when the gRNA and the off-target sequence have less than six mismatches, this can result in off-target consequences [84,85]. These effects can cause harmful outcomes, such as sequence mutations, deletions, and genome rearrangements, which hinder the application of CRISPR/Cas9 for certain objectives. Although there have been no documented instances of off-target consequences of this system in the diamondback moth, it is hypothesized that such occurrences may still happen and compromise the precision of the CRISPR/Cas9 system. In order to tackle this problem, several approaches may be utilized to improve the precision of the CRISPR/Cas9 system. These include refining the sgRNA sequence, altering the Cas9 gene, and employing alternative Cas variants. Cas variants, such as Cas12a and Cas13, offer unique advantages for genome editing and gene regulation. Cas12a (Cpf1) is an RNA-guided endonuclease that introduces staggered double-stranded breaks, producing cohesive ends that are advantageous for precise gene insertions [86,87]. Unlike Cas9, Cas12a recognizes T-rich PAM sequences, which expand its target range, particularly in AT-rich genomes [88]. Furthermore, it operates using a single crRNA, simplifying the design and delivery of editing components. Cas13, on the other hand, is unique in targeting RNA rather than DNA [89,90]. This makes it particularly useful for transient gene knockdown, studying RNA function, and combating RNA-based pathogens. Its high specificity for RNA reduces the risk of permanent off-target effects, making it a promising tool for applications requiring reversible gene regulation [89]. These variants, with their distinct mechanisms and features, could be helpful in overcoming the off-target challenges of CRISPR/Cas9 while broadening the scope of genetic interventions in P. xylostella.
Additionally, delivering CRISPR/Cas9 components into target organisms can be crucial, especially for in vivo applications, and may influence the efficiencies [91]. In the case of P. xylostella, the ribonucleoproteins have mainly been utilized as CRISPR/Cas9 components and delivered through embryo microinjections (Table 1). Microinjection to early embryos is a difficult, time-consuming, and laborious procedure. Furthermore, the post-injection survival rate of individuals varies and relies on expertise, which poses a challenge in making this system effective for P. xylostella (Table 1 and Table 2). Other delivery methods, including receptor-mediated ovary transduction of cargo (ReMOT) control, have been successfully developed for mosquitoes and B. mori. In this method, adult individuals and pupae were injected with Cas9-mediated ribonucleoprotein complexes, resulting in significant germline mutations [92,93]. This method can be optimized for P. xylostella, which may help to overcome challenges related to the delivery of components to the diamondback moth. However, its application to P. xylostella may present some potential limitations and risks. One major limitation would be the variability in delivery efficiency as the method relies on the compatibility of ligand–receptor interactions, which may not be fully characterized for P. xylostella. Additionally, unintended off-target effects could occur if the cargo interacts with non-target tissues or receptors, potentially leading to undesired physiological or developmental changes. Furthermore, the potential for incomplete or inconsistent germline editing may hinder its application in population-level genetic control strategies. Despite these challenges, further research into optimizing ligands and improving delivery specificity could enhance the feasibility of ReMOT control for P. xylostella and might offer advantages over traditional germline delivery methods.
The efficiency of gene-editing using the CRISPR/Cas9 system in the diamondback moth depends on the components used for microinjections. For instance, using a ribonucleoprotein complex to knockout the target gene results in a significantly better mutation rate compared to using messenger RNA (mRNA) with synthetic sgRNA and plasmid DNA (Table 2). It is crucial to acknowledge that the editing that occurs in somatic tissues cannot be passed on to the next generation. Only germline mutations can be inherited, making it crucial for the effectiveness of this system in pest management. In P. xylostella, high somatic cleavage efficiency, i.e., exceeding 70%, hinders the transmission of mutant alleles to offspring [69,70]. To address this challenge, it is essential for Cas9 to cause high cleavage at the target gene in germ cells rather than in somatic tissues. This can be accomplished by limiting the activity of Cas9 in germline cells, necessitating the use of highly specialized promoters that selectively activate Cas9 in germ tissues.
The main challenge of the CRISPR/Cas9 system is the low effectiveness of the homing-based gene drive and the formation of high-resistance alleles in the diamondback moth. The CRISPR/Cas9 gene drive operates primarily on the principles of HDR, which involve introducing exogenous gene sequences. This leads to the creation of DSBs at the target locus of an organism, followed by repair through HDR. However, in most cases, the cells mainly rely on NHEJ to repair these DSBs, leading to resistance allele formation. The NHEJ repair process causes the formation of alleles that are resistant to gene drive components. This leads to the elimination of the target site for the gRNA and hampers the propagation of gene drive components among populations. In our previous studies, Cas9 efficiently cleaves the target gene; however, the cleaved place is more often repaired through the NHEJ process rather than the HDR process, resulting in the formation of resistance alleles [69,70]. To overcome this challenge, two basic strategies can be employed: inhibiting the NHEJ pathway to promote HDR-mediated repair, and activating the HDR pathway. There are specific genes involved in the NHEJ-mediated DNA repair pathway, such as DNA-dependent protein kinase (DNA-PK), Ku80, Ku70, and ligase 4 (LIG4). DNA-PK acts as an initial component in recognizing and binding DSBs during NHEJ repair; the Ku70/80 heterodimer plays an important role for initiating the NHEJ pathway by showing a strong affinity for dsDNA ends; and LIG4 is a vital enzyme that controls the NHEJ repair process. Inhibiting DNA-PK, Ku70/80, and LIG4 significantly enhances the HDR-mediated repair mechanism in various organisms, including humans, mice, and insects (B. mori) [94,95,96,97,98,99,100,101]. Conversely, HDR is a crucial pathway for the precise repair of DSBs. An alternative strategy to shift DNA repair from NHEJ to HDR involves inducing the activation of key components within the HDR machinery [102,103]. The CtIP protein is crucial in the early phases of HDR, and the integration of transgenes by HDR can be improved by combining Cas9 with the N-terminal domain of CtIP [104]. MLN4924 increases CtIP expression levels by inhibiting neddylation, thereby effectively enhancing HDR efficiency [105]. Rad51 plays a crucial role in facilitating DNA strand exchange during HDR. Heterologous expression of Rad51 has been shown to boost HDR efficiency [106]. The G2/M phase of the cell cycle is another critical window during which HDR predominantly occurs. Utilizing small molecules capable of reversibly arresting cells at this stage significantly improves overall HDR efficiency [107,108,109,110].
Beyond improving HDR efficiency, alternative genome-editing strategies offer potential solutions to mitigate resistance allele formation. Prime editing is emerging as a promising approach, enabling precise genetic modification without inducing DSBs [111,112]. Additionally, advanced techniques such as PASTE (programmable addition via site-specific targeting elements) and TwinPE (twin prime editing) facilitate large DNA insertions with high precision, reducing the risk of resistance allele formation [113]. These techniques could be integrated into gene drive systems to enhance their efficiency and stability. Combining HDR enhancement strategies with prime editing and its derivatives could provide a robust framework for improving gene drive efficiency in P. xylostella. Future research should explore the feasibility of integrating these advanced genome-editing tools to achieve more stable and effective genetic control strategies while minimizing resistance allele formation. By focusing on these pathways, the challenges related to low HDR efficiency can be addressed in the future, making this system more effective for genetic manipulations in P. xylostella.

5. Conclusions and Future Perspectives

This review highlights the remarkable progress and applications of CRISPR/Cas9 genome editing in the diamondback moth (P. xylostella), a destructive pest responsible for substantial agricultural losses. This technology has facilitated precise genetic modifications, providing critical insights into the molecular mechanisms underlying reproduction, development, insecticide resistance, and host adaptability. Such advancements lay the foundation for innovative, genetic-based pest control strategies that could significantly reduce reliance on chemical insecticides, thereby promoting sustainable agricultural practices for P. xylostella. Despite its considerable potential, the widespread application of CRISPR/Cas9 in pest management faces several technical and ecological challenges. Off-target mutations, inefficient delivery methods, variable knockout efficiencies, low germline editing, and low HDR rates remain key obstacles to be addressed. Additionally, concerns regarding unintended ecological consequences, resistance evolution, and the ethical implications of gene drives necessitate stringent risk assessments and regulatory oversight.
Future research should focus on optimizing HDR-mediated repair, enhancing germline transmission, and exploring alternative CRISPR variants such as CRISPR/Cas12 and CRISPR/Cas13 to improve genome editing in P. xylostella. Furthermore, integrating CRISPR-based strategies with ecological approaches, such as integrated pest management (IPM) and biological control, will enhance their sustainability and long-term efficacy. By aligning CRISPR/Cas9 applications with ecological and regulatory considerations, this technology holds great promise for revolutionizing pest management. Through these efforts, the full potential of CRISPR/Cas9 in pest management can be achieved, leading to more sustainable and effective solutions for controlling diamondback moth.

Author Contributions

M.A. and G.Y. contributed to the conceptualization and design of this study; Y.C. and J.L. were responsible for writing the original draft and data visualization; G.Y. and M.A. performed the final review and editing. All authors have read and agreed to the published version of the manuscript.

Funding

This work was funded by the Key Project of Fujian Province Natural Science Foundation (grant number: 2023J02009); the Fuzhou–Xiamen–Quanzhou National Independent Innovation Demonstration Zone RNAi Microbial Pesticide Collaborative Innovation Platform Project; the Fujian Province Central Government-Guided Local Science and Technology Development Fund Project (grant number: 2024L3004); and the “111” program (grant number: KRA16001A).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Li, Z.; Feng, X.; Liu, S.-S.; You, M.; Furlong, M.J. Biology, ecology, and management of the diamondback moth in China. Annu. Rev. Entomol. 2016, 61, 277–296. [Google Scholar] [CrossRef] [PubMed]
  2. You, M.; Ke, F.; You, S.; Wu, Z.; Liu, Q.; He, W.; Baxter, S.W.; Yuchi, Z.; Vasseur, L.; Gurr, G.M. Variation among 532 genomes unveils the origin and evolutionary history of a global insect herbivore. Nat. Commun. 2020, 11, 2321. [Google Scholar] [CrossRef] [PubMed]
  3. Dancau, T.; Mason, P.G.; Cappuccino, N. Elusively overwintering: A review of diamondback moth (Lepidoptera: Plutellidae) cold tolerance and overwintering strategy. Can. Entomol. 2018, 150, 156–173. [Google Scholar] [CrossRef]
  4. Sun, J.; Liang, P.; Gao, X. Cross-resistance patterns and fitness in fufenozide-resistant diamondback moth, Plutella xylostella (Lepidoptera: Plutellidae). Pest Manag. Sci. 2012, 68, 285–289. [Google Scholar] [CrossRef]
  5. Etebari, K.; Furlong, M.J.; Asgari, S. Genome wide discovery of long intergenic non-coding RNAs in Diamondback moth (Plutella xylostella) and their expression in insecticide resistant strains. Sci. Rep. 2015, 5, 14642. [Google Scholar] [CrossRef]
  6. Harvey-Samuel, T.; Ant, T.; Alphey, L. Towards the genetic control of invasive species. Biol. Invasions 2017, 19, 1683–1703. [Google Scholar] [CrossRef]
  7. Kyrou, K.; Hammond, A.M.; Galizi, R.; Kranjc, N.; Burt, A.; Beaghton, A.K.; Nolan, T.; Crisanti, A. A CRISPR-Cas9 gene drive targeting doublesex causes complete population suppression in caged Anopheles gambiae mosquitoes. Nat. Biotechnol. 2018, 36, 1062–1066. [Google Scholar] [CrossRef]
  8. Asad, M.; Liu, D.; Chen, J.; Yang, G. Applications of gene drive systems for population suppression of insect pests. Bull. Entomol. Res. 2022, 112, 724–733. [Google Scholar] [CrossRef]
  9. Kandul, N.P.; Liu, J.; Buchman, A.; Shriner, I.C.; Corder, R.M.; Warsinger-Pepe, N.; Yang, T.; Yadav, A.K.; Scott, M.J.; Marshall, J.M. Precision guided sterile males suppress populations of an invasive crop pest. GEN Biotechnol. 2022, 1, 372–385. [Google Scholar] [CrossRef]
  10. Li, M.; Yang, T.; Bui, M.; Gamez, S.; Wise, T.; Kandul, N.P.; Liu, J.; Alcantara, L.; Lee, H.; Edula, J.R.; et al. Suppressing mosquito populations with precision guided sterile males. Nat. Commun. 2021, 12, 5374. [Google Scholar] [CrossRef]
  11. Ishino, Y.; Shinagawa, H.; Makino, K.; Amemura, M.; Nakata, A. Nucleotide sequence of the iap gene, responsible for alkaline phosphatase isozyme conversion in Escherichia coli, and identification of the gene product. J. Bacteriol. 1987, 169, 5429–5433. [Google Scholar] [CrossRef] [PubMed]
  12. Jansen, R.; Embden, J.D.A.v.; Gaastra, W.; Schouls, L.M. Identification of genes that are associated with DNA repeats in prokaryotes. Mol. Microbiol. 2002, 43, 1565–1575. [Google Scholar] [CrossRef] [PubMed]
  13. Gostimskaya, I. CRISPR-Cas9: A history of its discovery and ethical considerations of its use in genome editing. Biochemistry 2022, 87, 777–788. [Google Scholar] [CrossRef] [PubMed]
  14. Mali, P.; Yang, L.; Esvelt, K.M.; Aach, J.; Guell, M.; DiCarlo, J.E.; Norville, J.E.; Church, G.M. RNA-guided human genome engineering via Cas9. Science 2013, 339, 823–826. [Google Scholar] [CrossRef]
  15. Cong, L.; Ran, F.A.; Cox, D.; Lin, S.; Barretto, R.; Habib, N.; Hsu, P.D.; Wu, X.; Jiang, W.; Marraffini, L.A. Multiplex genome engineering using CRISPR/Cas systems. Science 2013, 339, 819–823. [Google Scholar] [CrossRef]
  16. Hwang, W.Y.; Fu, Y.; Reyon, D.; Maeder, M.L.; Tsai, S.Q.; Sander, J.D.; Peterson, R.T.; Yeh, J.R.J.; Joung, J.K. Efficient genome editing in zebrafish using a CRISPR-Cas system. Nat. Biotechnol. 2013, 31, 227–229. [Google Scholar] [CrossRef]
  17. Hille, F.; Charpentier, E. CRISPR-Cas: Biology, mechanisms and relevance. Philos. Trans. R. Soc. B Biol. Sci. 2016, 371, 20150496. [Google Scholar] [CrossRef]
  18. Xue, C.; Greene, E.C. DNA repair pathway choices in CRISPR-Cas9-mediated genome editing. Trends Genet. 2021, 37, 639–656. [Google Scholar] [CrossRef]
  19. Wang, J.Y.; Doudna, J.A. CRISPR technology: A decade of genome editing is only the beginning. Science 2023, 379, eadd8643. [Google Scholar] [CrossRef]
  20. Hillary, V.E.; Ceasar, S.A. A Review on the Mechanism and Applications of CRISPR/Cas9/Cas12/Cas13/Cas14 Proteins Utilized for Genome Engineering. Mol. Biotechnol. 2023, 65, 311–325. [Google Scholar] [CrossRef]
  21. Jinek, M.; Chylinski, K.; Fonfara, I.; Hauer, M.; Doudna, J.A.; Charpentier, E. A programmable dual-RNA–guided DNA endonuclease in adaptive bacterial immunity. Science 2012, 337, 816–821. [Google Scholar] [CrossRef] [PubMed]
  22. Asmamaw, M.; Zawdie, B. Mechanism and applications of CRISPR/Cas-9-mediated genome editing. Biol. Targets Ther. 2021, 15, 353–361. [Google Scholar] [CrossRef]
  23. Ceasar, S.A.; Maharajan, T.; Hillary, V.E.; Krishna, T.P.A. Insights to improve the plant nutrient transport by CRISPR/Cas system. Biotechnol. Adv. 2022, 59, 107963. [Google Scholar] [CrossRef] [PubMed]
  24. Liu, R.; Chen, L.; Jiang, Y.; Zhou, Z.; Zou, G. Efficient genome editing in filamentous fungus Trichoderma reesei using the CRISPR/Cas9 system. Cell Discov. 2015, 1, 15007. [Google Scholar] [CrossRef]
  25. Lee, H.; Yoon, D.E.; Kim, K. Genome editing methods in animal models. Anim. Cells Syst. 2020, 24, 8–16. [Google Scholar] [CrossRef]
  26. Singh, V.; Dhar, P.K. Genome Engineering via CRISPR-Cas9 System; Academic Press: Cambridge, MA, USA, 2020. [Google Scholar]
  27. Hillary, V.E.; Ceasar, S.A. Genome engineering in insects for the control of vector borne diseases. Prog. Mol. Biol. Transl. Sci. 2021, 179, 197–223. [Google Scholar]
  28. Garza-Elizondo, M.A.; Rodríguez-Rodríguez, D.R.; Ramírez-Solís, R.; Garza-Rodríguez, M.D.L.; Barrera-Saldaña, H.A. Genome editing: A perspective on the application of CRISPR/Cas9 to study human diseases. Int. J. Mol. Med. 2019, 43, 1559–1574. [Google Scholar]
  29. Meccariello, A.; Monti, S.M.; Romanelli, A.; Colonna, R.; Primo, P.; Inghilterra, M.G.; Del Corsano, G.; Ramaglia, A.; Iazzetti, G.; Chiarore, A.; et al. Highly efficient DNA-free gene disruption in the agricultural pest Ceratitis capitata by CRISPR-Cas9 ribonucleoprotein complexes. Sci. Rep. 2017, 7, 10061. [Google Scholar] [CrossRef]
  30. Huang, Y.; Chen, Y.; Zeng, B.; Wang, Y.; James, A.A.; Gurr, G.M.; Yang, G.; Lin, X.; Huang, Y.; You, M. CRISPR/Cas9 mediated knockout of the abdominal-A homeotic gene in the global pest, diamondback moth (Plutella xylostella). Insect Biochem. Mol. Biol. 2016, 75, 98–106. [Google Scholar] [CrossRef]
  31. Zhao, Q.; Ma, D.; Huang, Y.; He, W.; Li, Y.; Vasseur, L.; You, M. Genome-wide investigation of transcription factors provides insights into transcriptional regulation in Plutella xylostella. Mol. Genet. Genomics 2017, 293, 435–449. [Google Scholar] [CrossRef]
  32. Zou, M.-M.; Wang, Q.; Chu, L.-N.; Vasseur, L.; Zhai, Y.-L.; Qin, Y.-D.; He, W.-Y.; Yang, G.; Zhou, Y.-Y.; Peng, L.; et al. CRISPR/Cas9-induced vitellogenin knockout lead to incomplete embryonic development in Plutella xylostella. Insect Biochem. Mol. Biol. 2020, 123, 103406. [Google Scholar] [CrossRef] [PubMed]
  33. Peng, L.; Wang, Q.; Zou, M.-M.; Qin, Y.-D.; Vasseur, L.; Chu, L.-N.; Zhai, Y.-L.; Dong, S.-J.; Liu, L.-L.; He, W.-Y.; et al. CRISPR/Cas9-mediated vitellogenin receptor knockout leads to functional deficiency in the reproductive development of Plutella xylostella. Front. Physiol. 2020, 10, 01585. [Google Scholar] [CrossRef] [PubMed]
  34. Xu, X.; Wang, Y.; Bi, H.; Xu, J.; Liu, Z.; Niu, C.; He, L.; James, A.A.; Li, K.; Huang, Y. Mutation of the seminal protease gene, serine protease 2, results in male sterility in diverse lepidopterans. Insect Biochem. Mol. Biol. 2020, 116, 103243. [Google Scholar] [CrossRef]
  35. Zhai, Y.-L.; Dong, S.-J.; Zou, M.-M.; Qin, Y.-D.; Liu, L.-L.; Cao, M.-H.; Huang, M.-Q.; Vasseur, L.; You, M.-S.; Peng, L. Vitelline membrane protein 26 mutagenesis, using CRISPR/Cas9, results in egg collapse in Plutella xylostella. Int. J. Mol. Sci. 2022, 23, 9538. [Google Scholar] [CrossRef]
  36. Liu, D.; Asad, M.; Liao, J.; Chen, J.; Li, J.; Chu, X.; Pang, S.; Tariq, M.; Abbas, A.N.; Yang, G. The potential role of the piwi gene in the development and reproduction of Plutella xylostella. Int. J. Mol. Sci. 2023, 24, 12321. [Google Scholar] [CrossRef]
  37. Zheng, W.; Ma, H.; Liu, Z.; Zhou, Y.; Zhu, H.; Liu, J.; Zhang, C.; Liu, Z.; Zhou, X. Knockout of tyramine receptor 1 results in a decrease of oviposition, mating, and sex pheromone biosynthesis in female Plutella xylostella. Pest Manag. Sci. 2023, 79, 3903–3912. [Google Scholar] [CrossRef]
  38. Asad, M.; Liao, J.; Chen, J.; Munir, F.; Pang, S.; Abbas, A.N.; Yang, G. Exploring the role of the ovary-serine protease gene in the female fertility of the diamondback moth using CRISPR/Cas9. Pest Manag. Sci. 2024, 80, 3194–3206. [Google Scholar] [CrossRef]
  39. Huang, P.; Yu, H.; Asad, M.; Liao, J.; Lin, S.; Pang, S.; Chu, X.; Yang, G. Functional characteristics of Dicer genes in Plutella xylostella. Pest Manag. Sci. 2024, 80, 2109–2119. [Google Scholar] [CrossRef]
  40. Xu, X.; Harvey-Samuel, T.; Yang, J.; Alphey, L.; You, M. Ommochrome pathway genes kynurenine 3-hydroxylase and cardinal participate in eye pigmentation in Plutella xylostella. BMC Mol. Cell Biol. 2020, 21, 63. [Google Scholar] [CrossRef]
  41. Xu, X.; Harvey-Samuel, T.; Yang, J.; You, M.; Alphey, L. CRISPR/Cas9-based functional characterization of the pigmentation gene ebony in Plutella xylostella. Insect Mol. Biol. 2021, 30, 615–623. [Google Scholar] [CrossRef]
  42. Wang, Y.; Huang, Y.; Xu, X.; Liu, Z.; Li, J.; Zhan, X.; Yang, G.; You, M.; You, S. CRISPR/Cas9-based functional analysis of yellow gene in the diamondback moth, Plutella xylostella. Insect Sci. 2020, 28, 1504–1509. [Google Scholar] [CrossRef] [PubMed]
  43. Wang, Y.; Chen, X.e.; Liu, Z.; Xu, J.; Li, X.; Bi, H.; Andongma, A.A.; Niu, C.; Huang, Y. Mutation of doublesex induces sex-specific sterility of the diamondback moth Plutella xylostella. Insect Biochem. Mol. Biol. 2019, 112, 103180. [Google Scholar] [CrossRef] [PubMed]
  44. Wang, Y.; Xu, X.; Chen, X.e.; Li, X.; Bi, H.; Xu, J.; Zhu, C.; Niu, C.; Huang, Y. Mutation of P-element somatic inhibitor induces male sterility in the diamondback moth, Plutella xylostella. Pest Manag. Sci. 2021, 77, 3588–3596. [Google Scholar] [CrossRef] [PubMed]
  45. Cao, M.H.; Zou, M.M.; Liu, L.L.; Dong, S.J.; Huang, M.Q.; Zheng, J.H.; Li, R.N.; Cui, J.D.; Peng, L. Sast1-mediated manifold effects inhibit Plutella xylostella fertility. Pest Manag. Sci. 2024, 80, 2596–2609. [Google Scholar] [CrossRef] [PubMed]
  46. Chen, S.P.; Wang, D.F.; Ma, W.F.; Lin, X.L.; Yang, G. Knockout of cryptochrome 1 disturbs the locomotor circadian rhythm and development of Plutella xylostella. Insect Sci. 2022, 30, 1035–1045. [Google Scholar] [CrossRef]
  47. Chen, S.-P.; Lin, X.-L.; Qiu, R.-Z.; Chi, M.-X.; Yang, G. An lw-opsin mutation changes the gene expression of the phototransduction pathway: A cryptochrome1 mutation enhances the phototaxis of male Plutella xylostella (lepidoptera: Plutellidae). Insects 2023, 14, 72. [Google Scholar] [CrossRef]
  48. Chen, W.; Wang, D.; Yu, L.; Zhong, W.; Yuan, Y.; Yang, G. Comparative analysis of locomotor behavior and head diurnal transcriptome regulation by PERIOD and CRY2 in the diamondback moth. Insect Sci. 2024, 31, 1697–1720. [Google Scholar] [CrossRef]
  49. Chen, W.; Dong, Y.; Saqib, H.S.A.; Vasseur, L.; Zhou, W.; Zheng, L.; Lai, Y.; Ma, X.; Lin, L.; Xu, X.; et al. Functions of duplicated glucosinolate sulfatases in the development and host adaptation of Plutella xylostella. Insect Biochem. Mol. Biol. 2020, 119, 103316. [Google Scholar] [CrossRef]
  50. Chen, W.; Dong, Y.; Zheng, L.; Lai, Y.; Li, F.; Zhou, L.; Wang, B.; You, M.; He, W. An inducible gene from glycoside hydrolase one family of Plutella xylostella decreases larval survival when feeding on host plant. Front. Physiol. 2022, 13, 1013092. [Google Scholar] [CrossRef]
  51. Liu, X.-L.; Zhang, J.; Yan, Q.; Miao, C.-L.; Han, W.-K.; Hou, W.; Yang, K.; Hansson, B.S.; Peng, Y.-C.; Guo, J.-M.; et al. The molecular basis of host selection in a crucifer-specialized moth. Curr. Biol. 2020, 30, 4476–4482.e4475. [Google Scholar] [CrossRef]
  52. Chen, W.; Saqib, H.S.A.; Xu, X.; Dong, Y.; Zheng, L.; Lai, Y.; Jing, X.; Lu, Z.; Sun, L.; You, M.; et al. Glucosinolate sulfatases–sulfatase-modifying factors system enables a crucifer-specialized moth to pre-detoxify defensive glucosinolate of the host plant. J. Agric. Food Chem. 2022, 70, 11179–11191. [Google Scholar] [CrossRef] [PubMed]
  53. Wang, B.-B.; Lai, Y.-F.; Li, F.-F.; Jiao, L.; Qiao, Q.-X.; Li, S.-Y.; Xiang, X.-J.; Liao, H.; You, M.-S.; He, W.-Y. Functional Characterization of Two RNA Methyltransferase Genes METTL3 and METTL14 Uncovers the Roles of m6A in Mediating Adaptation of Plutella xylostella to Host Plants. Int. J. Mol. Sci. 2022, 23, 10013. [Google Scholar] [CrossRef] [PubMed]
  54. Lei, G.; Zhou, H.; Chen, Y.; Vasseur, L.; Gurr, G.M.; You, M.; You, S. A very long-chain fatty acid enzyme gene, PxHacd2 affects the temperature adaptability of a cosmopolitan insect by altering epidermal permeability. Sci. Total Environ. 2023, 891, 164372. [Google Scholar] [CrossRef]
  55. Zhou, H.; Lei, G.; Chen, Y.; You, M.; You, S. Pxtret1-like affects the temperature adaptability of a cosmopolitan pest by altering trehalose tissue distribution. Int. J. Mol. Sci. 2022, 23, 9019. [Google Scholar] [CrossRef] [PubMed]
  56. Liu, Y.; Zhang, S.; Cao, S.; Jacquin-Joly, E.; Zhou, Q.; Liu, Y.; Wang, G. An odorant receptor mediates the avoidance of Plutella xylostella against parasitoid. BMC Biol. 2024, 22, 61. [Google Scholar] [CrossRef]
  57. Lei, G.; Huang, J.; Zhou, H.; Chen, Y.; Song, J.; Xie, X.; Vasseur, L.; You, M.; You, S. Polygenic adaptation of a cosmopolitan pest to a novel thermal environment. Insect Mol. Biol. 2024, 33, 387–404. [Google Scholar] [CrossRef]
  58. Guest, M.; Goodchild, J.A.; Bristow, J.A.; Flemming, A.J. RDL A301S alone does not confer high levels of resistance to cyclodiene organochlorine or phenyl pyrazole insecticides in Plutella xylostella. Pestic. Biochem. Physiol. 2019, 158, 32–39. [Google Scholar] [CrossRef]
  59. Guo, Z.; Sun, D.; Kang, S.; Zhou, J.; Gong, L.; Qin, J.; Guo, L.; Zhu, L.; Bai, Y.; Luo, L.; et al. CRISPR/Cas9-mediated knockout of both the PxABCC2 and PxABCC3 genes confers high-level resistance to Bacillus thuringiensis Cry1Ac toxin in the diamondback moth, Plutella xylostella (L.). Insect Biochem. Mol. Biol. 2019, 107, 31–38. [Google Scholar] [CrossRef]
  60. Sun, D.; Zhu, L.; Guo, L.; Wang, S.; Wu, Q.; Crickmore, N.; Zhou, X.; Bravo, A.; Soberón, M.; Guo, Z.; et al. A versatile contribution of both aminopeptidases N and ABC transporters to Bt Cry1Ac toxicity in the diamondback moth. BMC Biol. 2022, 20, 33. [Google Scholar] [CrossRef]
  61. Xiong, L.; Liu, Z.; Li, J.; Yao, S.; Li, Z.; Chen, X.; Shen, L.; Zhang, Z.; Li, Y.; Hou, Q.; et al. Analysis of the effect of Plutella xylostella polycalin and ABCC2 transporter on Cry1Ac susceptibility by CRISPR/Cas9-mediated knockout. Toxins 2023, 15, 273. [Google Scholar] [CrossRef]
  62. Zhao, S.; Jiang, D.; Wang, F.; Yang, Y.; Tabashnik, B.E.; Wu, Y. Independent and synergistic effects of knocking out two ABC transporter genes on resistance to Bacillus thuringiensis toxins Cry1Ac and Cry1Fa in diamondback moth. Toxins 2020, 13, 9. [Google Scholar] [CrossRef] [PubMed]
  63. Wang, X.; Ma, Y.; Wang, F.; Yang, Y.; Wu, S.; Wu, Y. Disruption of nicotinic acetylcholine receptor α6 mediated by CRISPR/Cas9 confers resistance to spinosyns in Plutella xylostella. Pest Manag. Sci. 2019, 76, 1618–1625. [Google Scholar] [CrossRef] [PubMed]
  64. Wang, X.; Cao, X.; Jiang, D.; Yang, Y.; Wu, Y. CRISPR/Cas9 mediated ryanodine receptor I4790M knockin confers unequal resistance to diamides in Plutella xylostella. Insect Biochem. Mol. Biol. 2020, 125, 103453. [Google Scholar] [CrossRef]
  65. Ye, M.; Xiong, L.; Dong, Y.; Xie, C.; Zhang, Z.; Shen, L.; Li, Z.; Yue, Z.; Jiang, P.; Yuchi, Z.; et al. The potential role of the methionine aminopeptidase gene pxmetap1 in a cosmopolitan pest for Bacillus thuringiensis toxin tolerance. Int. J. Mol. Sci. 2022, 23, 13005. [Google Scholar] [CrossRef]
  66. Sun, X.; Hua, W.; Wang, K.; Song, J.; Zhu, B.; Gao, X.; Liang, P. A novel V263I mutation in the glutamate-gated chloride channel of Plutella xylostella (L.) confers a high level of resistance to abamectin. Int. J. Biol. Macromol. 2023, 230, 123389. [Google Scholar] [CrossRef]
  67. Jiang, D.; Yu, Z.; He, Y.; Wang, F.; Gu, Y.; Davies, T.G.E.; Fan, Z.; Wang, X.; Wu, Y. Key role of the ryanodine receptor I4790K mutation in mediating diamide resistance in Plutella xylostella. Insect Biochem. Mol. Biol. 2024, 168, 104107. [Google Scholar] [CrossRef]
  68. You, S.; Yao, S.; Chen, X.; Hou, Q.; Liu, Z.; Lei, G.; Xie, X.; Liang, Z.; Yuchi, Z.; You, M.; et al. CRISPR/Cas9-mediated knockout of the PxJHBP gene resulted in increased susceptibility to Bt Cry1Ac protoxin and reduced lifespan and spawning rates in Plutella xylostella. J. Agric. Food Chem. 2024, 72, 8180–8188. [Google Scholar] [CrossRef]
  69. Asad, M.; Liu, D.; Li, J.; Chen, J.; Yang, G. Development of CRISPR/Cas9-mediated gene-drive construct targeting the phenotypic gene in Putella xylostella. Front. Physiol. 2022, 13, 938621. [Google Scholar] [CrossRef]
  70. Xu, X.; Harvey-Samuel, T.; Siddiqui, H.A.; Ang, J.X.D.; Anderson, M.E.; Reitmayer, C.M.; Lovett, E.; Leftwich, P.T.; You, M.; Alphey, L. Toward a CRISPR-Cas9-based gene drive in the diamondback moth Plutella xylostella. CRISPR J. 2022, 5, 224–236. [Google Scholar] [CrossRef]
  71. Chaves, I.; Pokorny, R.; Byrdin, M.; Hoang, N.; Ritz, T.; Brettel, K.; Essen, L.O.; van der Horst, G.T.J.; Batschauer, A.; Ahmad, M. The cryptochromes: Blue light photoreceptors in plants and animals. Annu. Rev. Plant Biol. 2011, 62, 335–364. [Google Scholar] [CrossRef]
  72. Young, M.W.; Kay, S.A. Time zones: A comparative genetics of circadian clocks. Nat. Rev. Genet. 2001, 2, 702–715. [Google Scholar] [CrossRef] [PubMed]
  73. Fahey, J.W.; Zalcmann, A.T.; Talalay, P. The chemical diversity and distribution of glucosinolates and isothiocyanates among plants. Phytochemistry 2002, 59, 237. [Google Scholar] [CrossRef]
  74. Halkier, B.A.; Gershenzon, J. Biology and biochemistry of glucosinolates. Annu. Rev. Plant Biol. 2006, 57, 303–333. [Google Scholar] [CrossRef]
  75. Chen, W.; Amir, M.B.; Liao, Y.; Yu, H.; He, W.; Lu, Z. New insights into the plutella xylostella detoxifying enzymes: Sequence evolution, structural similarity, functional diversity, and application prospects of glucosinolate sulfatases. J. Agric. Food Chem. 2023, 71, 10952–10969. [Google Scholar] [CrossRef] [PubMed]
  76. Li, B.J.; Wang, K.K.; Chen, D.P.; Yan, Y.; Cai, X.L.; Chen, H.M.; Dong, K.; Lin, F.; Xu, H.H. Distinct roles of two RDL GABA receptors in fipronil action in the diamondback moth (Plutella xylostella). Insect Sci. 2021, 28, 1721–1733. [Google Scholar] [CrossRef]
  77. Anderson, M.A.E.; Gonzalez, E.; Edgington, M.P.; Ang, J.X.D.; Purusothaman, D.-K.; Shackleford, L.; Nevard, K.; Verkuijl, S.A.N.; Harvey-Samuel, T.; Leftwich, P.T.; et al. A multiplexed, confinable CRISPR/Cas9 gene drive can propagate in caged Aedes aegypti populations. Nat. Commun. 2024, 15, 729. [Google Scholar] [CrossRef]
  78. Sinkins, S.P.; Gould, F. Gene drive systems for insect disease vectors. Nat. Rev. Genet. 2006, 7, 427–435. [Google Scholar] [CrossRef]
  79. Werren, J.H. Selfish genetic elements, genetic conflict, and evolutionary innovation. Proc. Nat. Acad. Sci. USA 2011, 108, 10863–10870. [Google Scholar] [CrossRef]
  80. Grunwald, H.A.; Gantz, V.M.; Poplawski, G.; Xu, X.-R.S.; Bier, E.; Cooper, K.L. Super-Mendelian inheritance mediated by CRISPR–Cas9 in the female mouse germline. Nature 2019, 566, 105–109. [Google Scholar] [CrossRef]
  81. Hammond, A.; Galizi, R.; Kyrou, K.; Simoni, A.; Siniscalchi, C.; Katsanos, D.; Gribble, M.; Baker, D.; Marois, E.; Russell, S.; et al. A CRISPR-Cas9 gene drive system targeting female reproduction in the malaria mosquito vector Anopheles gambiae. Nat. Biotechnol. 2016, 34, 78–83. [Google Scholar] [CrossRef]
  82. Zhang, Z.; Xiong, L.; Xie, C.; Shen, L.; Chen, X.; Ye, M.; Sun, L.; Yang, X.; Yao, S.; Yue, Z.; et al. Optimization and application of CRISPR/Cas9 genome editing in a cosmopolitan pest, diamondback moth. Int. J. Mol. Sci. 2022, 23, 13042. [Google Scholar] [CrossRef] [PubMed]
  83. Li, S.; Lin, G.; Wen, H.; Lu, H.; Yin, A.; Zheng, C.; Li, F.; Qiao, Q.; Jiao, L.; Lin, L.; et al. Knock-in of exogenous sequences based on CRISPR/Cas9 targeting autosomal genes and sex chromosomes in the diamondback moth, Plutella xylostella. J. Integr. Agric. 2024, 23, 3089–3103. [Google Scholar] [CrossRef]
  84. Zhang, X.H.; Tee, L.Y.; Wang, X.G.; Huang, Q.S.; Yang, S.H. Off-target effects in CRISPR/Cas9-mediated genome engineering. Mol. Ther. Nucleic Acids 2015, 4, e264. [Google Scholar] [CrossRef]
  85. Kaeuferle, T.; Stief, T.A.; Canzar, S.; Kutlu, N.N.; Willier, S.; Stenger, D.; Ferrada-Ernst, P.; Habjan, N.; Peters, A.E.; Busch, D.H.; et al. Genome-wide off-target analyses of CRISPR/Cas9-mediated T-cell receptor engineering in primary human T cells. Clin. Transl. Immun. 2022, 11, e1372. [Google Scholar] [CrossRef]
  86. Safari, F.; Zare, K.; Negahdaripour, M.; Barekati-Mowahed, M.; Ghasemi, Y. CRISPR Cpf1 proteins: Structure, function and implications for genome editing. Cell Biosci. 2019, 9, 36. [Google Scholar] [CrossRef]
  87. Kim, H.; Lee, W.-j.; Oh, Y.; Kang, S.-H.; Hur, J.K.; Lee, H.; Song, W.; Lim, K.-S.; Park, Y.-H.; Song, B.-S.; et al. Enhancement of target specificity of CRISPR–Cas12a by using a chimeric DNA–RNA guide. Nucleic Acids Res. 2020, 48, 8601–8616. [Google Scholar] [CrossRef]
  88. Swarts, D.C.; Jinek, M. Cas9 versus Cas12a/Cpf1: Structure–function comparisons and implications for genome editing. WIREs RNA 2018, 9, e1481. [Google Scholar] [CrossRef]
  89. Kordyś, M.; Sen, R.; Warkocki, Z. Applications of the versatile CRISPR-Cas13 RNA targeting system. WIREs RNA 2022, 13, e1694. [Google Scholar] [CrossRef]
  90. Yang, H.; Patel, D.J. Structures, mechanisms and applications of RNA-centric CRISPR–Cas13. Nat. Chem. Biol. 2024, 20, 673–688. [Google Scholar] [CrossRef]
  91. Zhang, S.; Shen, J.; Li, D.; Cheng, Y. Strategies in the delivery of Cas9 ribonucleoprotein for CRISPR/Cas9 genome editing. Theranostics 2021, 11, 614–648. [Google Scholar] [CrossRef]
  92. Chaverra-Rodriguez, D.; Macias, V.M.; Hughes, G.L.; Pujhari, S.; Suzuki, Y.; Peterson, D.R.; Kim, D.; McKeand, S.; Rasgon, J.L. Targeted delivery of CRISPR-Cas9 ribonucleoprotein into arthropod ovaries for heritable germline gene editing. Nat. Commun. 2018, 9, 3008. [Google Scholar] [CrossRef] [PubMed]
  93. Yu, B.; Dong, S.; Jiang, X.; Qiao, L.; Chen, J.; Li, T.; Pan, G.; Zhou, Z.; Li, C. Cas9-mediated gene editing using receptor-mediated ovary transduction of cargo (ReMOT) control in Bombyx mori. Insects 2023, 14, 923. [Google Scholar] [CrossRef] [PubMed]
  94. Riesenberg, S.; Maricic, T. Targeting repair pathways with small molecules increases precise genome editing in pluripotent stem cells. Nat. Commun. 2018, 9, 2164. [Google Scholar] [CrossRef] [PubMed]
  95. Tavecchio, M.; Munck, J.M.; Cano, C.; Newell, D.R.; Curtin, N.J. Further characterisation of the cellular activity of the DNA-PK inhibitor, NU7441, reveals potential cross-talk with homologous recombination. Cancer Chemother. Pharmacol. 2012, 69, 155–164. [Google Scholar] [CrossRef]
  96. Chu, V.T.; Weber, T.; Wefers, B.; Wurst, W.; Sander, S.; Rajewsky, K.; Kühn, R. Increasing the efficiency of homology-directed repair for CRISPR-Cas9-induced precise gene editing in mammalian cells. Nat. Biotechnol. 2015, 33, 543–548. [Google Scholar] [CrossRef]
  97. Li, G.; Liu, D.; Zhang, X.; Quan, R.; Zhong, C.; Mo, J.; Huang, Y.; Wang, H.; Ruan, X.; Xu, Z.; et al. Suppressing Ku70/Ku80 expression elevates homology-directed repair efficiency in primary fibroblasts. Int. J. Biochem. Cell Biol. 2018, 99, 154–160. [Google Scholar] [CrossRef]
  98. Hu, Z.; Shi, Z.; Guo, X.; Jiang, B.; Wang, G.; Luo, D.; Chen, Y.; Zhu, Y.-S. Ligase IV inhibitor SCR7 enhances gene editing directed by CRISPR–Cas9 and ssODN in human cancer cells. Cell Biosci. 2018, 8, 12. [Google Scholar] [CrossRef]
  99. Maruyama, T.; Dougan, S.K.; Truttmann, M.C.; Bilate, A.M.; Ingram, J.R.; Ploegh, H.L. Increasing the efficiency of precise genome editing with CRISPR-Cas9 by inhibition of nonhomologous end joining. Nat. Biotechnol. 2015, 33, 538–542. [Google Scholar] [CrossRef]
  100. Yao, X.; Zhang, M.L.; Wang, X.; Ying, W.Q.; Hu, X.D.; Dai, P.F.; Meng, F.L.; Shi, L.Y.; Sun, Y.; Yao, N.; et al. Tild-CRISPR Allows for Efficient and Precise Gene Knockin in Mouse and Human Cells. Dev. Cell 2018, 45, 526–536.e5. [Google Scholar] [CrossRef]
  101. Zhu, L.; Mon, H.; Xu, J.; Lee, J.M.; Kusakabe, T. CRISPR/Cas9-mediated knockout of factors in non-homologous end joining pathway enhances gene targeting in silkworm cells. Sci. Rep. 2015, 5, 18103. [Google Scholar] [CrossRef]
  102. Sartori, A.A.; Lukas, C.; Coates, J.; Mistrik, M.; Fu, S.; Bartek, J.; Baer, R.; Lukas, J.; Jackson, S.P. Human CtIP promotes DNA end resection. Nature 2007, 450, 509–514. [Google Scholar] [CrossRef] [PubMed]
  103. Anand, R.; Ranjha, L.; Cannavo, E.; Cejka, P. Phosphorylated CTIP functions as a co-factor of the mre11-rad50-nbs1 endonuclease in DNA end resection. Mol. Cell 2016, 64, 940–950. [Google Scholar] [CrossRef] [PubMed]
  104. Charpentier, M.; Khedher, A.H.Y.; Menoret, S.; Brion, A.; Lamribet, K.; Dardillac, E.; Boix, C.; Perrouault, L.; Tesson, L.; Geny, S.; et al. CtIP fusion to Cas9 enhances transgene integration by homology-dependent repair. Nat. Commun. 2018, 9, 1133. [Google Scholar] [CrossRef]
  105. Jimeno, S.; Fernández-Ávila, M.J.; Cruz-García, A.; Cepeda-García, C.; Gómez-Cabello, D.; Huertas, P. Neddylation inhibits CtIP-mediated resection and regulates DNA double strand break repair pathway choice. Nucleic Acids Res. 2015, 43, 987–999. [Google Scholar] [CrossRef]
  106. Yu, S.; Song, Z.; Luo, J.; Dai, Y.; Li, N. Over-expression of RAD51 or RAD54 but not RAD51/4 enhances extra-chromosomal homologous recombination in the human sarcoma (HT-1080) cell line. J. Biotechnol. 2011, 154, 21–24. [Google Scholar] [CrossRef]
  107. Cooper, S.; Iyer, G.; Tarquini, M.; Bissett, P. Nocodazole does not synchronize cells: Implications for cell-cycle control and whole-culture synchronization. Cell Tissue Res. 2006, 324, 237–242. [Google Scholar] [CrossRef]
  108. Hande, K.R.; Hagey, A.; Berlin, J.; Cai, Y.; Meek, K.; Kobayashi, H.; Lockhart, A.C.; Medina, D.; Sosman, J.; Gordon, G.B.; et al. The pharmacokinetics and safety of ABT-751, a novel, orally bioavailable sulfonamide antimitotic agent: Results of a phase 1 study. Clin. Cancer Res. 2006, 12, 2834–2840. [Google Scholar] [CrossRef]
  109. Rahman, S.H.; Bobis-Wozowicz, S.; Chatterjee, D.; Gellhaus, K.; Pars, K.; Heilbronn, R.; Jacobs, R.; Cathomen, T. The nontoxic cell cycle modulator indirubin augments transduction of adeno-associated viral vectors and zinc-finger nuclease-mediated gene targeting. Hum. Gene Ther. 2013, 24, 67–77. [Google Scholar] [CrossRef]
  110. Lin, S.; Staahl, B.T.; Alla, R.K.; Doudna, J.A. Enhanced homology-directed human genome engineering by controlled timing of CRISPR/Cas9 delivery. eLife 2014, 3, e04766. [Google Scholar] [CrossRef]
  111. Chen, P.J.; Liu, D.R. Prime editing for precise and highly versatile genome manipulation. Nat. Rev. Genet. 2023, 24, 161–177. [Google Scholar] [CrossRef]
  112. Zeng, H.; Daniel, T.C.; Lingineni, A.; Chee, K.; Talloo, K.; Gao, X. Recent advances in prime editing technologies and their promises for therapeutic applications. Curr. Opin. Biotechnol. 2024, 86, 103071. [Google Scholar] [CrossRef]
  113. Mikkelsen, N.S.; Bak, R.O. Enrichment strategies to enhance genome editing. J. Biomed. Sci. 2023, 30, 51. [Google Scholar] [CrossRef] [PubMed]
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Figure 1. Life cycle of P. xylostella. The life cycle of P. xylostella (from hatching to adulthood) typically lasts 18–25 days, varying based on temperature and geographic location. The larval stage (from the first to the fourth instar) is the longest phase of the diamondback moth’s life cycle.
Figure 1. Life cycle of P. xylostella. The life cycle of P. xylostella (from hatching to adulthood) typically lasts 18–25 days, varying based on temperature and geographic location. The larval stage (from the first to the fourth instar) is the longest phase of the diamondback moth’s life cycle.
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Figure 2. Schematic illustration of the CRISPR/Cas9 system in the diamondback moth. Two basic methods of the CRISPR/Cas9 genome-editing system are being used in the diamondback moth. (A) The knockout method: a mixture of ribonucleoprotein (gRNA + Cas9) is injected into the eggs of the diamondback moth. The gRNA binds to a specific target gene, and Cas9 creates a DSB. The cleaved DNA repairs through NHEJ and directs the knockout of the particular target gene. (B) The knock-in method: a mixture of ribonucleoprotein and donor plasmid is injected into the eggs of the diamondback moth. The gRNA binds to the target gene and Cas9 creates a DSB. The cleaved DNA repairs through HDR and copies the donor DNA fragment to the cleaved target site.
Figure 2. Schematic illustration of the CRISPR/Cas9 system in the diamondback moth. Two basic methods of the CRISPR/Cas9 genome-editing system are being used in the diamondback moth. (A) The knockout method: a mixture of ribonucleoprotein (gRNA + Cas9) is injected into the eggs of the diamondback moth. The gRNA binds to a specific target gene, and Cas9 creates a DSB. The cleaved DNA repairs through NHEJ and directs the knockout of the particular target gene. (B) The knock-in method: a mixture of ribonucleoprotein and donor plasmid is injected into the eggs of the diamondback moth. The gRNA binds to the target gene and Cas9 creates a DSB. The cleaved DNA repairs through HDR and copies the donor DNA fragment to the cleaved target site.
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Figure 3. Schematic illustration of gene drive systems. (A) depicts the global gene drive where Cas9, gRNA, and a marker gene are inserted into the target locus. Under normal conditions, Cas9 cleaves the target locus in germ cells and converts the normal (wild-type) allele into a gene drive allele that bypasses Mendelian segregation. (B) illustrates the split-drive system where Cas9 and one marker gene are inserted into an autonomous chromosome, while gRNA and another marker gene are inserted into the target locus. Two independent transgenic lines are created and crossed. The drive allele is inherited more frequently than the normal Mendelian segregation due to the active presence of Cas9 and gRNA in germline cells. This schematic representation illustrates germline drive conversion, highlighting the process of germline homing.
Figure 3. Schematic illustration of gene drive systems. (A) depicts the global gene drive where Cas9, gRNA, and a marker gene are inserted into the target locus. Under normal conditions, Cas9 cleaves the target locus in germ cells and converts the normal (wild-type) allele into a gene drive allele that bypasses Mendelian segregation. (B) illustrates the split-drive system where Cas9 and one marker gene are inserted into an autonomous chromosome, while gRNA and another marker gene are inserted into the target locus. Two independent transgenic lines are created and crossed. The drive allele is inherited more frequently than the normal Mendelian segregation due to the active presence of Cas9 and gRNA in germline cells. This schematic representation illustrates germline drive conversion, highlighting the process of germline homing.
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Table 1. Utilization of CRISPR/Cas9-based system in the diamondback moth.
Table 1. Utilization of CRISPR/Cas9-based system in the diamondback moth.
ApplicationsGene Knockout/
Knock-In		CRISPR/Cas9 ComponentMethod of DeliveryGene Function and Disruption ImpactReferences
Development and reproductionPxHomeoboxKnockoutCas9 mRNAEmbryo injectionsTranscription factor; lethal to embryos[31]
PxVgKnockoutRibonucleoproteinEmbryo injectionsEgg maturation; reduced hatching rate[32]
PxVgRKnockoutCas9 mRNAEmbryo injectionsOocyte development; smaller eggs; low fertility[33]
PxSer2KnockoutRibonucleoproteinEmbryo injectionsSemen protein; male sterility[34]
PxVMPKnockoutRibonucleoproteinEmbryo injectionsEgg structure formation; egg collapse and reduced hatching[35]
PxPiwiKnockoutRibonucleoproteinEmbryo injectionsGerm cell development; delayed pupation; failed emergence[36]
PxTAR1KnockoutRibonucleoproteinEmbryo injectionsGerm cell development; delayed pupation,;failed emergence[37]
PxOspKnockoutRibonucleoproteinEmbryo injectionsOvarian development; fewer and irregular eggs[38]
PxDcr-1KnockoutRibonucleoproteinEmbryo injectionsRNA processing; increased mortality; reduced reproduction[39]
PxDcr-2KnockoutCas9 mRNAEmbryo injectionsRNA processing; reduced RNAi efficiency[39]
PigmentationPxKMOKnockoutRibonucleoproteinEmbryo injectionsEye pigment gene; yellow/red eye color[40]
PxCardinalKnockoutRibonucleoproteinEmbryo injectionsEye pigment gene; yellow eyes turning red with age[40]
PxEbonyKnockoutRibonucleoproteinEmbryo injectionsDopamine metabolism; darker body color[41]
PxYellowKnockoutRibonucleoproteinEmbryo injectionsBody pigmentation gene; change in body color[42]
Sex-determinationPxDsxKnockoutCas9 mRNAEmbryo injectionsSex differentiation; genital abnormalities[43]
PxPSIKnockoutCas9 mRNAEmbryo injectionsMale-specific splicing; genital defects[44]
PxSast1KnockoutRibonucleoproteinEmbryo injectionsSex differentiation; male infertility; egg defects[45]
Circadian rhythmsPxCry1KnockoutRibonucleoproteinEmbryo injectionsLight-sensitive photoreceptor; altered activity rhythms[46]
PxLW-opsinKnockoutRibonucleoproteinEmbryo injectionsPhototaxis; impaired light response[47]
PxCry2KnockoutRibonucleoproteinEmbryo injectionsCircadian rhythm; disturb the rhythmic activities[48]
PxPerKnockoutRibonucleoproteinEmbryo injectionsCircadian rhythm; activity disruption[48]
Ecological adaptabilityPxGSS1KnockoutRibonucleoproteinEmbryo injectionsGlucosinolate metabolism; reduced host plant adaptation[49,50]
PxGSS2KnockoutRibonucleoproteinEmbryo injectionsGlucosinolate metabolism; reduced host plant adaptation[49,50]
PxOr35KnockoutRibonucleoproteinEmbryo injectionsOlfactory receptor; oviposition preference reduced[51]
PXOr49KnockoutRibonucleoproteinEmbryo injectionsOlfactory receptor; oviposition preference reduced[51]
PxGSS3KnockoutRibonucleoproteinEmbryo injectionsGlucosinolate metabolism; host adaptability unchanged[52]
PxMETTL14KnockoutRibonucleoproteinEmbryo injectionsRNA methyltransferase; developmental defects[53]
Px008848(PxGH1)KnockoutRibonucleoproteinEmbryo injectionsInsect–plant interaction; increased larval survival[50]
PxHacd2KnockoutRibonucleoproteinEmbryo injectionsEnzyme for VLCFAs synthesis; survival and fecundity decreased [54]
PxTret1-likeKnockoutRibonucleoproteinEmbryo injectionsTrehalose transport; reduced temperature resistance[55]
PxOr16KnockoutRibonucleoproteinEmbryo injectionsOdor receptor; reduced avoidance behavior to parasitoid[56]
PxGCC2KnockoutRibonucleoproteinEmbryo injectionsPeripheral gene for stress; reduced stress resistance[57]
PxKPNB1KnockoutRibonucleoproteinEmbryo injectionsPeripheral gene for stress; decreased environmental adaptation[57]
Insecticide resistancePxGABAR-α1 (A282S)Knock-inCas9 protein crRNA, tracRNA and ssODNAEmbryo injectionsGABA-gated chloride channel; not involved in pesticide resistance[58]
PxABCC2KnockoutRibonucleoprotein/Cas9 mRNAEmbryo injectionsBt resistance; reduced Cry1Ac sensitivity[51,59,60,61,62]
PxABCC3KnockoutRibonucleoprotein/Cas9 mRNAEmbryo injectionsBt resistance; reduced Cry1Ac sensitivity[51,59,60,62]
PxnAChRα6KnockoutRibonucleoproteinEmbryo injectionsNicotinic acetylcholine receptor α6; increased pesticide resistance[63]
PxRyR (I4790M)Knock-inRibonucleoprotein and ssODNEmbryo injectionsRyanodine receptor; increased diamine resistance”[64]
PxAPN1KnockoutRibonucleoproteinEmbryo injectionsAminopeptidases increased Cry1Ac resistance[60]
PxAPN3aKnockoutRibonucleoproteinEmbryo injectionsAminopeptidases increased Cry1Ac resistance[60]
PxMetAP1KnockoutRibonucleoproteinEmbryo injectionsMethionine aminopeptidases; impaired peptide processing[65]
PxGluCl (V263I)Knock-inCas9 proteinss/sgRNA/ssODNEmbryo injectionsGlutamate-gated chloride chann; resistance to avermectin[66]
PxPolycalinKnockoutRibonucleoproteinEmbryo injectionsBt toxin receptor; decreased Cry1Ac sensitivity[61]
PxRyR (I4790K)Knock-inRibonucleoprotein and ssODNEmbryo injectionsRyanodine receptor; chlorantraniliprole resistance[67]
PxJHBPKnockoutRibonucleoproteinEmbryo injectionsJH signaling regulator; increased Cry1Ac susceptibility[68]
Gene drivePxYellowKnock-inRibonucleoprotein and plasmidEmbryo injectionsBody pigmentation gene; change in body color[69,70]
PxKmoKnock-inRibonucleoprotein and plasmidEmbryo injectionsEye pigment gene; yellow/red eye color[70]
Table 2. Efficiency of CRISPR/Cas9 system in the diamondback moth.
Table 2. Efficiency of CRISPR/Cas9 system in the diamondback moth.
Target GeneNo. of Eggs InjectedSurvived IndividualsSurvival RateMutation RateReference
PxABCC22158037.21%65.00%[59]
PxABCC32006834.00%58.82%[59]
PxDsxC48915130.87%62.91%[43]
PxDsxF43112629.23%34.13%[43]
PxDsxM51810720.66%42.99%[43]
PxGSS12055225.36%3.85%[52]
PxGSS22055225.36%5.77%[52]
PxVgR25812247.29%63.11%[33]
PxYellow67648071.01%56.66%[42]
PxSer2200049036.5%1.02%[34]
PxKmo2229442.3417.02%[40]
PxCardinal1825530.22%29.09%[40]
PxVg1359973.33%4.04%[32]
PxPSI63832450.78%23.15%[44]
PxCry12635822.05%20.69%[46]
PxMETTL142563011.72%10.00%[53]
PxVMP521019.23%22.22%[35]
PxTret1-like1006262%11.29%[55]
PxLW-opsin2116229.38%6.45%[47]
PxHacd21005656.00%3.57%[54]
PxPiwi2129846.23%55.10%[36]
PxGluCl (V263I)300113.67%9.09%[66]
PxTAR11546844.16%13.24%[37]
PxOsp18012368.33%19.25%[38]
PxSast11856836.80%1.47%[45]
PxPer2107234.29%4.17%[48]
PxRyR (I4790K)1350594.37%4.35%[67]
PxGCC21765330.11%5.66%[57]
PxKPNB11544227.27%9.52%[57]
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Asad, M.; Chang, Y.; Liao, J.; Yang, G. CRISPR/Cas9 Genome Editing in the Diamondback Moth: Current Progress, Challenges, and Prospects. Int. J. Mol. Sci. 2025, 26, 1515. https://doi.org/10.3390/ijms26041515

AMA Style

Asad M, Chang Y, Liao J, Yang G. CRISPR/Cas9 Genome Editing in the Diamondback Moth: Current Progress, Challenges, and Prospects. International Journal of Molecular Sciences. 2025; 26(4):1515. https://doi.org/10.3390/ijms26041515

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Asad, Muhammad, Yanpeng Chang, Jianying Liao, and Guang Yang. 2025. "CRISPR/Cas9 Genome Editing in the Diamondback Moth: Current Progress, Challenges, and Prospects" International Journal of Molecular Sciences 26, no. 4: 1515. https://doi.org/10.3390/ijms26041515

APA Style

Asad, M., Chang, Y., Liao, J., & Yang, G. (2025). CRISPR/Cas9 Genome Editing in the Diamondback Moth: Current Progress, Challenges, and Prospects. International Journal of Molecular Sciences, 26(4), 1515. https://doi.org/10.3390/ijms26041515

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