Next Article in Journal
Seasonal Dynamics and Factors Shaping Aquatic Insect Assemblages in Mountain Streams of the Pannonian Lowland Ecoregion
Previous Article in Journal
Developmental Regulation of Corazonin, Eclosion Hormone, and Bursicon Messages and RNAi Suppression of Corazonin in Adult, Female American Dog Ticks, Dermacentor variabilis
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Insects
Volume 16
Issue 4
10.3390/insects16040345
insects-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

CRISPR/Cas Technology in Insect Insecticide Resistance

by
Qiuchen Xu
†,
Mingyun Wang
†,
Jiahui Zeng
,
Hangzhen Sun
,
Xiaoqi Wei
,
Hui Jiang
,
Xuping Shentu
* and
Dan Sun
*
Key Laboratory of Microbiological Metrology, Measurement & Bio-Product Quality Security, State Administration for Market Regulation, College of Life Science, China Jiliang University, Hangzhou 310018, China
*
Authors to whom correspondence should be addressed.
†
These authors contributed equally to this work.
Insects 2025, 16(4), 345; https://doi.org/10.3390/insects16040345
Submission received: 28 November 2024 / Revised: 18 January 2025 / Accepted: 21 January 2025 / Published: 26 March 2025
(This article belongs to the Section Insect Pest and Vector Management)
Browse Figures
Review Reports Versions Notes

Simple Summary

Chemical and biological insecticides are crucial in modern agricultural, forestry, and public health pest management, and are involved in protecting crops, preserving human health, and maintaining ecological balance. However, the misuse of insecticides has led to increased insect resistance, mainly due to altered expression or mutations in insect target genes that encode proteins involved in pesticide recognition, binding, and detoxification. The emergence of CRISPR/Cas gene-editing technology has deepened our understanding of the mechanisms by which insects adapt to and resist insecticides. This article summarizes the progress of this technology in the prevention and control of agricultural pests and public health pests in different arthropods.

Abstract

Chemicals and biological insecticides play a crucial role as pest management strategies in modern agriculture and forestry. However, their excessive and unreasonable use inevitably leads to varying degrees of resistance among insect populations, which seriously affects the sustainability of insecticide use. One primary reason for this resistance is alterations or mutations in insect gene expression. One class of genes encodes proteins that serve as critical targets for insecticides to exert their toxic effects in insects, while another class of genes encodes proteins involved in the detoxification process of insecticides within insects. Reverse genetics has become a vital research tool for studying the molecular mechanisms underlying changes and mutations in these target genes and their impact on insect resistance. The advent of CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats) and the CRISPR-associated gene Cas as gene-editing technologies has significantly advanced our understanding of how insects adapt to and resist insecticides. This article aims to provide a comprehensive and objective review of the progress made using the CRISPR/Cas system in various arthropods within the field of pest control.

1. Introduction

Insects are an important part of biodiversity. They provide important ecosystem services such as pollination and pest control but also cause damage as disease vectors and plant pests [1,2,3]. Scientists have attempted to tackle these key issues through biological and chemical methods. However, extensive and widespread use of insecticides has led to pest resistance as a consequence, which impedes pest control effectiveness [4,5,6]. As the global population continues to rise, there will be a huge demand for food in the near future [7]. Crop pests are considered a serious challenge to agriculture production and threaten food security [8]. According to incomplete statistics, global annual crop losses caused by insect pest damage are up to hundreds of billions of dollars [9]. Public health pests often serve as vectors or carriers that transmit various bacteria and viruses posing threats to human health [10,11,12]. Biological and chemical insecticides serve as the two primary pest management strategies in agriculture, safeguarding crop yield and quality. However, the unscientific and non-standard application of these pesticides also causes increasing pest resistance [13,14,15]. The evolution of insect resistance to insecticides is frequently related to the expression alterations or mutations in the target site [16,17,18,19]. Certain key genes in insects exhibit altered expression levels upon exposure to pesticides, resulting in significant changes in the concentration of corresponding intracellular protein products [20]. These proteins often serve as primary targets for insecticides, such as Acetylcholinesterase (AChE) that serves as the primary target for commercial pesticides, particularly organophosphate and carbamate insecticides [21]. In many insects that develop resistance to pesticides, such as the DDT-resistant malaria vector Anopheles gambiae, the activity of Glutathione S-transferases (GSTs) increases significantly. This enhanced activity plays a crucial role in the resistance mechanism, enabling these insects to more effectively eliminate or neutralize pesticide molecules that enter their bodies [22]. For example, the high expression of CYP6F1 in Culex pipiens pallens can confer resistance to deltametharin [23]. Gene mutations can alter the amino acid sequence at the interaction sites between proteins and insecticides, resulting in target modifications that typically diminish insecticide efficacy and produce exceptionally high levels of resistance [24,25].
The rapid advancement of gene-editing technologies has facilitated studies that explore and manipulate specific gene functions in insects [26]. The first prevalent technology for targeted genome editing used zinc finger nucleases (ZFNs) [27]. ZFNs consist of zinc finger proteins that specifically recognize DNA sequences fused with Fok I nucleases. This setup induces double-strand breaks (DSBs) in DNA, activating the cellular DNA damage repair mechanisms and triggering gene mutations (Figure 1A) [28,29]. As early as 2002, ZFNs were applied to insect species. For instance, Bibikova M and colleagues mutated the color gene in Drosophila using ZFNs [30]. Beumer K and others produced mutated offspring of fruit flies through the same method [31]. Subsequently, ZFNs were also applied in other insect models, such as Caenorhabditis elegans [32] and Danaus plexippus [28]. Although ZFNs played a pivotal role in early insect gene-editing research, their relatively complex preparation process and high costs have largely limited their widespread application. Subsequently, transcription-activator-like effector nucleases (TALENs) were developed, which achieved the specific recognition and cleavage of DNA sequences with the help of Fok I nucleases (Figure 1B). TALEN exhibits higher specificity and lower toxicity than ZFNs when cutting the same sequence [33,34]. Scientists innovatively applied an enhanced TALEN technology to achieve rapid and efficient modifications of the Drosophila genome. They utilized the “modular assembly” strategy to combine smaller, recognizable DNA sequence units into larger, more complex protein structures capable of recognizing and cleaving specific DNA sequences, thereby constructing specific TALENs targeted at the yellow gene and a novel autosomal gene. Subsequent mRNA injections into Drosophila embryos yielded results indicating that 31.2% of the fertile F0 generation carried heritable modifications of the yellow gene, completing the entire process in just one month [35].
The CRISPR/Cas genetic modification platform represents a groundbreaking advancement in gene editing, fostering novel opportunities for the creation of biopesticides and the investigation of pesticide resistance mechanisms [36]. The CRISPR/Cas system is currently in widespread use, including various types such as CRISPR/Cas12a (also known as Cpf1), CRISPR/Cas13a, and CRISPR/Cas13b, each with unique advantages and application potentials. Among the many members of the CRISPR/Cas system, CRISPR/Cas9 stands out as the most prominent tool. Its efficiency, precision, and ease of use have quickly made it the mainstream tool in the field of gene editing. Notably, the key difference between CRISPR/Cas9 and ZFN and TALEN lies in its use of short RNA sequences as specificity recognition units, activating the Cas9 enzyme to induce double-strand breaks in target DNA [37] (Figure 1C). Whilst, compared with ZFNs and TALENs, CRISPR/Cas has demonstrated significant advantages in the study of insecticide resistance. Its high efficiency, simplicity, low cost, and ease of use enable researchers to quickly and accurately cut and edit target genes, thereby precisely manipulating genes related to insecticide resistance in a short period of time, observing the effects of genetic changes on the phenotype of organisms, and revealing their roles and mechanisms in drug resistance [38,39,40,41,42]. Moreover, the changes at the genomic level produced by CRISPR/Cas are stable and heritable, allowing mutated genes to transfer to the next generation [43,44,45].
The CRISPR/Cas technology has garnered significant attention in the field of entomology, largely due to the successful genomic editing of the classic model organism Drosophila melanogaster [46]. Subsequently, CRISPR/Cas has been effectively utilized in various insects, including Plutella xylostella [47], An. gambiae [48,49], and Bombyx mori [50]. The rapid advancements in omics and molecular biology in recent years have led to an increasing application of CRISPR/Cas in entomological studies. However, there is currently a lack of a comprehensive review article that systematically explores and integrates the application of CRISPR/Cas technology in insect insecticide resistance. This article reviews the application and prospects of the CRISPR/Cas system concerning insecticide resistance in Diptera, Lepidoptera, and Hemiptera. The emergence of CRISPR/Cas technology presents new strategies and methodologies for exploring insect gene functions and field pest control. As technology continues to advance and improve, CRISPR/Cas-based pest management strategies will play a more crucial role in agricultural production, contributing significantly to food security, ecological safety, and human health.

2. Applications of CRISPR/Cas9 in Insecticide Resistance

The studies that have used CRISPR/Cas9 in insecticide resistance have been generalized according to species from different insect orders and the types of gene editing (Figure 2). Detailed information has also been organized to facilitate a comparison (Table 1).

2.1. Diptera

2.1.1. Drosophila melanogaster

As a classic model organism in biological research, D. melanogaster has consistently been at the forefront of genetic analysis, significantly contributing to the advancement of entomology [114]. The CRISPR/Cas9 technology serves as a crucial tool in understanding various genotypes, receptors, and ion channels that influence resistance formation in Drosophila. CYP genes are pivotal members of insect metabolic detoxifying enzymes, playing a vital role in the development of insect resistance. In D. melanogaster, CYP6G1 and CYP6G2 are key candidate genes involved in metabolism and resistance to multiple insecticides. Denecke utilized CRISPR/Cas9 to knockout CYP6G1 and CYP6G2 genes, revealing that the high expression of these two genes confers imidacloprid resistance [51]. Fusetto further demonstrated that CYP6G1 is associated with the formation of insecticide oxidative metabolites [52]. Insect cell membrane ion channels are pivotal targets for a wide array of insecticides, with functional diversity being intrinsically associated with pest resistance. The kdr mutation in voltage-gated sodium channels (vgscs) is notably linked to resistance to prevalent insecticides in insects. The kdr mutation was successfully introduced into D. melanogaster using CRISPR/Cas9 and subsequently evaluated the sensitivity variations in these mutants to permethrin DDT and deltamethrin. Furthermore, they employed CRISPR-based allele drives to substitute the resistant kdr mutations with their susceptible wild-type alleles. This foundational validation paves the way for numerous potential applications, such as the targeted reversion of insecticide-resistant populations to a state of susceptibility [53]. A combination of genetic transformation and CRISPR/Cas9 genome-editing technology demonstrated that the transgenic D. melanogaster strains expressing P450 enzymes along with engineered mutations in the voltage-gated sodium channel (para) are associated with pyrethroid resistance [54]. The nicotine acetylcholine receptor (nAChR), a neurotransmitter-gated ion channel protein, is widely distributed in the central nervous system of insects and serves as a critical target for various insecticides. The G275E mutation in the nAChR Dα6 subunit imparts resistance to spinosad, achieved through CRISPR/Cas9-mediated gene knock-in in D. melanogaster [55]. A systematic study of the nAChR gene family in D. melanogaster was conducted using CRISPR/Cas9 to explore the response mechanisms to three different classes of insecticides: neonicotinoids, spinosyns, and sulfoximines [56]. Recently, researchers developed a sensitivity gene drive at the acetylcholine esterase (Ace) locus in D. melanogaster using CRISPR/Cas9 technology, and they successfully replaced resistant mutation types with sensitive ones, potentially reversing pest resistance and restoring sensitivity to pesticides [57]. Similarly, CASPP ((5-chloro-1′-[(E)-3-(4-chlorophenyl)allyl] spiro[indoline-3,4′-piperidine]-1-yl}-(2-chloro-4-pyridyl)methanone) compounds represent a novel class of neuroactive insecticides that specifically target the Vesicular Acetylcholine Transporter (VAChT). Vernon integrated transgenic D. melanogaster lines with the GAL4/UAS system to enhance the transcription level of VAChT. In unison, they introduced the VAChTY49N mutation using CRISPR/Cas technology and found that D. melanogaster develop resistance to CASPP when the expression of wild-type VAChT is upregulated or there is a specific point mutation (VAChTY49N) [58]. ABC transporters play a pivotal role in insecticide resistance, facilitating the transport of various neuroactive insecticides. The knockdown of three genes, Mdr65, Mdr49, and Mdr50, of the ABC transporter in D. melanogaster using CRISPR/Cas9 revealed that the mutation of Mdr65 increased the sensitivity to a range of neuroactive insecticides, whereas the responses of Mdr49 and Mdr50 varied depending on the specific insecticide. The combination of the CRISPR/Cas9 method and ABC inhibitor verapamil further confirms the role of ABC transporters in enhancing insecticide tolerance in D. melanogaster and unveils their tissue and substrate specificity [59].
Insects with GABA receptors, particularly those composed of the RDL (resistant to dieldrin) subunit, are critical targets for commonly used synthetic insecticides. Zhou examined the sensitivities of seven distinct D. melanogaster RDL point mutants to four meta-diamide and isoxazoline insecticides [60]. CRISPR/Cas9 technology has significantly contributed to understanding insecticide resistance in D. melanogaster by modifying key genes such as CYPs, nAChR, VAChT, ABC transporters, and ion channels, providing insights into resistance mechanisms and facilitating the development of new pest management strategies.

2.1.2. Aedes aegypti, Culex quinquefasciatus, and Anopheles gambiae

Mosquitoes pose severe threats to human health by transmitting numerous diseases such as Zika, malaria, dengue, chikungunya, and filariasis [115,116,117]. These vector-borne viruses have emerged as a critical public health concern, affecting approximately 4 billion people worldwide. Vector control, including the use of insecticide-treated nets (ITNs) and indoor residual spraying (IRS), has proven effective in reducing malaria transmission in epidemic and hyperendemic areas. The application of insecticides is a crucial component of this initiative. However, the rise in insecticide resistance among mosquitoes and pathogens has led to the resurgence of these diseases [118].
Membrane proteins play a pivotal role in protecting cells from foreign substances, including drugs and toxins. Researchers have studied these proteins, particularly P-glycoprotein encoded by the MDR49 gene, and found that they confer resistance to certain insecticides, such as ivermectin, and can influence the reproductive capabilities of mosquitoes like Ae. aegypti [119]. Other membrane proteins, such as cadherin (Cad) and membrane-bound alkaline phosphatase (mALP), are potential receptors for Cry toxins from Bacillus thuringiensis, although the exact relationship between these proteins and Cry toxin activities remains unclear.
Using the CRISPR/Cas9 system, scientists have successfully generated knockout mutants of these membrane proteins in mosquitoes. For instance, Pacheco generated mALP- and Cad-knockout mutants in Ae. aegypti [61], while Lan identified ten candidate nicotinic acetylcholine receptor (nAChR) subunits and created a mutant strain lacking the Aaeα6 gene, which exhibited increased resistance to spinosad [120]. In addition to Ae. aegypti, C. quinquefasciatus is another global vector for various human and animal diseases [121,122]. Feng developed a specific Cas9/sgRNA expression toolkit for C. quinquefasciatus, providing significant support for the development of control technologies for this species [62]. Researchers have also used CRISPR genome-editing techniques to introduce mutations, such as the kdr mutation L1014F in An. gambiae, to investigate their effects on insecticide resistance [63]. Williams et al. confirmed that the V402L confers resistance to permethrin and DDT in An. Gambiae via the CRISPR/Cas9 system [64].
To prevent the transmission of mosquito-borne viruses, two primary strategies are proposed, i.e., reducing the capacity of vectors to harbor pathogens and suppressing insect populations through sterilization [123]. Membrane proteins and their mutations, studied using CRISPR/Cas9, play a crucial role in insecticide resistance and could be targets for developing new control measures against mosquito-borne diseases.

2.2. Lepidoptera

In recent years, CRISPR/Cas9 technology has introduced new tools for analyzing functional genes across various insect species. Among the Lepidoptera, B. mori is notable as the first agricultural insect to successfully undergo efficient genome editing using CRISPR/Cas9 for functional gene studies [26]. This order also encompasses several significant agricultural pests, such as P. xylostella, H. armigera, S. exigua, and S. frugiperda, which pose serious threats to crop yields. The emergence of resistance has rendered previously effective chemical control measures unreliable, complicating pest management considerably. By employing CRISPR/Cas technology to modify the specific genes within these pests, researchers can influence the growth, development, reproduction, and environmental adaptability of target insects, thus achieving pest control objectives.

2.2.1. Helicoverpa armigera, Helicoverpa zea, and Pectinophora gossypiella

H. armigera is a notorious global pest that infests cotton crops. Researchers have identified cadherin (CAD) as the receptor for the Bt Cry1A toxin in H. armigera. Wang employed CRISPR/Cas9 technology to successfully knockout the HaCad gene in a susceptible H. armigera population, establishing a homozygous SCD-Cad knockout strain that exhibited 549-fold resistance to the Cry1Ac toxin than that of the susceptible stain [85]. Subsequently, they also knocked out two Rdl homolog genes (HaRdl-1 and HaRdl-2), which encode critical subunits of insect γ-aminobutyric acid (GABA) receptors, the primary targets of cyclodiene and phenylpyrazole insecticides. The findings indicate that HaRdl-1 and HaRdl-2 play essential roles in determining the susceptibility of cotton bollworms to cyclodiene and other insecticides, exhibiting distinct pharmacological characteristics [86]. Furthermore, Wang discovered that the high resistance of H. armigera to the Bt toxin Cry2Ab is closely linked to a loss-of-function mutation in its ATP-binding cassette gene (ABCA2) [87]. In response, they developed two HaABCA2 knockout strains, designated as SCD-A2KO1 and SCD-A2KO2. The results reveal that both knockout strains display extreme resistance to Cry2Aa and Cry2Ab toxins while showing minimal or no resistance to Cry1Ac toxin. Further investigations demonstrate that the resistance of these knockout strains to Cry2Ab is recessive, with the resistance locus located on the HaABCA2 gene [88]. Zhang generated a series of homozygous knockout strains, including two single-gene knockout strains, in which the ATP-binding cassette (ABCC2) or CAD genes were deleted, and exhibited 512- or 396-fold Cry1Ac resistance ratio, respectively. Additionally, a double-gene knockout strain with HaABCC2 and HaCAD was simultaneously deleted and showed 6273-fold resistance to Cry1Ac toxin, significantly greater than single-gene knockouts and confirming the interaction of ABCC2 and Cad in H. armigera [89]. Cytochrome P450 plays a critical role in insecticide resistance in H. armigera. Wang used CRISPR/Cas9 genome-editing system to knockout a cluster of nine P450 genes, creating a homozygous mutant strain devoid of the CYP6AE cluster. This mutant strain exhibited reduced tolerance to esfenvalerate and indoxacarb. Furthermore, they confirmed that no single CYP6AE gene is essential for the survival of H. armigera. This finding significantly contributes to the future identification of candidate genes associated with insecticide resistance [124]. The Glutathione S-transferase (GST) gene family plays a significant role in insect detoxification mechanisms. Researchers found that the GST gene cluster in the North China strain of H. armigera underwent strong selection, indicating a significant resistance to lambda-cyhalothrin resistance. They knocked out a GST cluster using the CRISPR/Cas9 system, significantly increasing the susceptibility of H. armigera to lambda-cyhalothrin [90]. There are also cotton pests. Fabrick knocked out the ABCA2 gene in H. zea and P. gossypiella, respectively, resulting in mutated populations resistant to Bt toxin Cry2Ab [91,92]. Research on H. armigera and related cotton pests using CRISPR/Cas9 technology has identified key genes involved in insecticide resistance, providing insights into the mechanisms of resistance and guiding strategies for pest management.

2.2.2. Plutella xylostella, Trichoplusia ni, and Tuta absoluta

T. ni and P. xylostella are significant agricultural pests that cause serious damage to crops. Bt (Bacillus thuringiensis) can produce a series of insecticidal proteins for eliminating pests [125]. Guo provided the first in vivo reverse genetic evidence that ABCC2, ABCC3, APN1, APN3a and ABCC1 serve as midgut functional receptors for Bt Cry1 toxins in P. xylostella [65,66,67]. Subsequently, Sun investigated the contributions of two ABC transport proteins (ABCC2 and ABCC3) and two aminopeptidases N (APN1 and APN3a) towards Bt Cry1Ac toxin resistance. Their findings indicated that double-gene knockouts significantly boosted resistance, especially a quadruple-gene knockout, which resulted in exceptionally high resistance, suggesting functional redundancy with mutual complementation among these receptors [68]. Additionally, functional redundancy between ABC transporters (ABCC2 and ABCC3) and PxmALP also has been demonstrated by using CRISPR/Cas9-mediated multiple-gene knockout strategy [69]. The CRISPR/Cas9-induced knockout of non-receptor paralogs demonstrated that the increased expression of these genes is responsible for diminishing the fitness costs of Cry1Ac resistance [67]. Liu demonstrated that a mutation in either PxABCC2 or PxABCC3 alone is insufficient for high-level Cry1Ac resistance, necessitating mutations in both genes [70]. Zhao showed that dual-gene knockout of PxABCC2 and PxABCC3 significantly enhances resistance to Cry1Fa compared to single-gene knockout [71]. The CRISPR/Cas9-based mutation of PxABCB1 in P. xylostella exhibited a 63-fold resistance to Cry1Ac toxin compared to the parental susceptible strain [72]. Ye and his team validated the potential role of the methionine aminopeptidase gene PxMetAP1 in a cosmopolitan pest for Bt Cry1Ac toxin tolerance using CRISPR/Cas9 technology, which revealed a significant increase in susceptibility to Cry1Ac toxin [73]. Similar genes also exist in Trichoplusia ni, and Yang generated a homozygous strain of T. ni with frameshift mutations in ABCA1 and ABCA2 through CRISPR/Cas9 mutagenesis. Their findings indicate that the ABCA2 mutants were highly resistant to Cry2Ab, whereas mutations in ABCA1 do not exhibit the same level of resistance [97]. Ma also utilized CRISPR/Cas9 technology to validate the role of the resistance locus ABCC2 in conferring resistance to the Bt toxin Cry1Ac in Trichoplusia ni. Subsequent hybridization experiments revealed that the F1-generation Cry1Ac-resistant strains exhibited higher Cry1Ac resistance levels compared to the ABCC2-knockout CRISPR strains, suggesting the presence of additional mutations that may be associated with Cry1Ac resistance [98]. The CRISPR/Cas9-mediated knockout of the PxJHBP gene exhibited increased sensitivity to Cry1Ac toxin, reduced lifespan, and decreased fertility. This highlights the crucial role of PxJHBP in Cry1Ac resistance and female reproductive regulation, offering new insights into designing insecticides or interference strategies targeting insect reproductive systems [74]. Researchers used a high-efficiency CRISPR/Cas9 system to knockout SE2 (short interspersed nuclear element (SINE—named SE2)) in the promoter of MAP4K4 gene from the resistant P. xylostella strain, thereby altering the transcript levels of midgut genes linked with Cry1Ac resistance that showed an expression landscape similar to that of the susceptible P. xylostella strain [75]. The CRISPR/Cas9-mediated knockout of a midgut microRNA miR-8545 significantly increased the susceptibility of P. xylostella to Cry1Ac toxin [76]. The relationship between a mutation (I1056M/F) in the chitin synthase 1 (CHS1) gene of P. xylostella and benzoylureas (BPU) was verified in the D. melanogaster model using the CRISPR/Cas9 system [77]. Guo created a site-specific mutation of the m6A site in JH esterase (PxJHE) by the CRISPR/Cas9-mediated homology-directed repair (HDR) pathway in P. xylostella, resulting in a slight increase in the susceptibility to Cry1Ac protein [78].
There are many different genes that enhance the resistance of P. xylostella to different types of insecticides. Additionally, a reverse genetic approach was employed to confirm the association between the nAChR α6 subunit and spinosad resistance in P. xylostella [79]. Sun identified a novel valine-to-isoleucine mutation (V263I) in the glutamate-gated chloride channel (GluCl) in the field strain of P. xylostella, demonstrating varied resistance levels to abamectin. They successfully established a P. xylostella knock-in strain expressing the homozygous V263I mutation via CRISPR/Cas9, which exhibited a high resistance (106.3-fold) to abamectin but significantly reduced fecundity [80]. Similarly, CRISPR/Cas9-mediated D472N substitution in the Rdl1 GABAR of P. xylostella confers low resistance to abamectin and endosulfan, with no effect on other GABAR-targeting insecticides [81]. Meta-diamide insecticides combat Lepidopteran pests by activating insect ryanodine receptors (RyRs). Wang first demonstrated the functional role of the I4790M mutation in PxRyR concerning meta-diamide resistance [82]. Jiang introduced the I4790K mutation in PxRyR, transforming sensitive populations into resistant ones, indicating a strong correlation of this mutation with resistance to various insecticides, including diamides [83]. F1845Y and V1848I are two mutations in the IVS6 fragment of the P. xylostella and Tuta absoluta sodium channel domains. To confirm their roles in indoxacarb resistance, Samantsidis employed CRISPR/Cas9 technology to create homozygous mutants carrying F1845Y or V1848I in D. melanogaster. Results indicated that both mutations conferred moderate resistance to indoxacarb, while the F1845Y homozygous mutant exhibited significant resistance to metaflumizone [108]. Certain insecticides, such as chlorantraniliprole and other diamide-class insecticides, disrupt insect muscle contractions by activating or inhibiting ryanodine receptors (RyRs), ultimately leading to insect mortality. Consequently, RyRs emerge as primary targets for these insecticides. These mutations were replicated in D. melanogaster by CRISPR/Cas9 and conducted toxicity assessments. Their findings reveal that the G4946V mutation significantly enhances resistance to flubendiamide and chlorantraniliprole of T. absoluta, while the I4790M mutation predominantly affects resistance to flubendiamide. These findings not only substantiate the significance of RyR mutations in diamide resistance but also shed light on the intricate ways in which these mutations can alter the binding affinity of different diamide insecticides, providing a deeper understanding of the molecular basis of resistance in pests [109]. The critical role of the RyR gene in conferring resistance to diamide insecticides was emphasized by further studies, underscoring the intricate interplay of multiple mutations in the evolution of this resistance [84].
Pests like T. ni and P. xylostella damage crops significantly. Bt toxins from Bacillus thuringiensis are effective against these pests. CRISPR/Cas9 studies have revealed that multiple genes, such as ABCC2, ABCC3, APN1, APN3a, and others, play roles in Bt toxin resistance through functional redundancy. Mutations in these genes enhance resistance to toxins like Cry1Ac and Cry1Fa. Similar genes and mutations are also found in Trichoplusia ni. CRISPR/Cas9 has been used to validate the roles of these genes and mutations in resistance, and to study their interactions. Additionally, genes like PxJHBP, microRNAs, and chitin synthase are also involved in resistance and reproductive regulation.

2.2.3. Spodoptera frugiperda, Spodoptera exigua, and Spodoptera litura

S. frugiperda, S. exigua, and S. litura all belong to the family Noctuidae. S. frugiperda is one of the most damaging pests globally and has developed resistance to Cry proteins. The Bt vegetative insecticidal protein Vip3Aa serves as an effective substitute for Cry proteins in controlling several major pests that have not yet demonstrated practical resistance [126]. CRISPR/Cas is crucial for understanding how insecticides affect the physiological functions of pests. Based on the understanding of these toxins and the mode of action of insecticides, we can develop more sensitive biomarkers or monitoring tools for the early detection of drug resistance in pest populations and develop more precise management strategies [127]. Jin discovered that knocking out the transcription factor gene SfMyb reduced the susceptibility of S. frugiperda to Vip3Aa, and these results may facilitate advancements in the proactive monitoring and management of pest resistance to Vip3Aa [128]. Liu found that the knockout of the midgut-specific chitin synthase gene SfCHS2, mediated by a retrotransposon, resulted in a high level of resistance to Vip3Aa. This finding is vital for the monitoring and management of pest resistance to Vip3Aa in S. frugiperda [94]. Shi established a knockout strain of the nAChR α6 subunit in S. frugiperda using a CRISPR/Cas9 system with dual sgRNAs to induce a substantial deletion of a DNA fragment. Their results demonstrated that the disruption of Sfα6 leads to a remarked increase in resistance to spinosyns [95]. Xu discovered that CYP304F1 mutated by CRISPR/Cas9 technology significantly diminished the resistance to β-cypermethrin and chlorpyrifos in S. litura [93]. These research findings provide new perspectives and methods for basic research on pest resistance. Researchers can further explore the molecular mechanisms, genetic laws, and the impact of environmental factors on drug resistance, providing a more solid theoretical basis for subsequent monitoring and management work.
S. exigua is a globally distributed pest. Zuo successfully introduced the RyR G4946E mutation into S. exigua using CRISPR/Cas9 technology. The mutant exhibited resistance to chlorantraniliprole, cyantraniliprole, and flubendiamide at levels of 223-fold, 336-fold, and over 1000-fold, respectively [99]. The CRISPR/Cas9-mediated knockout of P-glycoprotein (P-gp) significantly increased the susceptibility of S. exigua to abamectin and emamectin benzoate (EB), but not to spinosad, chlorfenapyr, beta-cypermethrin, carbosulfan indoxacarb, chlorpyrifos, phoxim, diafenthiuron, chlorfluazuron, chlorantraniliprole, or two Bt toxins (Cry1Ca and Cry1Fa) [100]. Zuo established a homozygous population (Seα6-KO), demonstrating that this gene is linked to resistance against spinosad and spinetoram [101]. Furthermore, Zuo introduced the G275E mutation of nAChR α6 found in palm thrips and whiteflies into S. exigua to validate the role of the G275E mutation in nAChR α6 in spinosyn resistance [102]. Wang identified nine nAChR subunits in S. exigua, namely, nAChR α1-α7 and nAChR β1-β2, and created a deletion mutant (Seα1-KO) using CRISPR/Cas9 for the Seα1 gene. Compared to the wild type, Seα1-KO exhibited significantly enhanced resistance to trifluoropyrimidine, dimehypo, and dinotefuran, while sensitivity to other insecticides remained unchanged [103]. Shi discovered significant yet differing enrichment of the CYP9A gene cluster between S. exigua and S. frugiperda through comparative genomics, which included species-specific duplications and sequence variations. CRISPR/Cas9 knockout experiments demonstrated that CYP9A contributes to the detoxification of plant defense compounds and various chemical insecticides. Additionally, the CYP9A gene cluster was closely associated with high pyrethroid resistance in field populations [96].

2.2.4. Ostrinia furnacalis and Chilo suppressalis

O. furnacalis is a significant pest affecting crops such as corn throughout China, from Heilongjiang to Hainan. This species has developed a high level of resistance to the insecticidal crystals of Bt, particularly the Cry proteins Cry1A and Cry1F. This resistance is associated with mutations in the ATP-binding cassette (ABC) transporter subfamily C gene, ABCC2. Wang employed the CRISPR/Cas9 technique to successfully create a homozygous strain of O. furnacalis featuring an 8 bp deletion mutation in the ABCC2 gene (OfC2-KO). Functional studies demonstrated a causal relationship between the truncated mutation of OfABCC2 and increased resistance to the Cry1Fa toxin [104]. Furthermore, Jin confirmed that OfCad mediates Cry1Ac toxicity through both in vitro and in vivo experiments. Compared to the wild type, the OfCad knockout strain exhibited moderate resistance to Cry1Ac while maintaining sensitivity to other Cry proteins [129]. Additional studies targeting the ABC transporter subfamily G gene, ABCG4, revealed that mutations in this gene resulted in increased resistance to Cry1 toxins and affected larval development and population parameters [105]. The resistance of the C. suppressalis to tetraniliprole is increasing, and RyR double mutations (I4758M and Y4667C) provide greater resistance to chlorfenapyr than single mutations [106]. Further evidence has shown that, in a single RyR mutation, Y4667D contributes the most to resistance to tetraphenylproline, followed by G4915E. Y4667C and I4758M are roughly equal, while Y4891F contributes the least [107]. This knowledge will support the development of innovative pest management strategies to address the escalating issue of resistance.

2.3. Hemiptera

Hemipteran species possess specialized piercing–sucking mouthparts that enable them to ingest liquid food. In herbivorous hemipterans, these intricate mouthparts facilitate the efficient extraction of nutrients from the xylem or phloem of plants. This adaptive evolution has led to certain hemipterans becoming significant pests in global agriculture, resulting in substantial annual crop losses. The long-term dependence on chemical pesticides in agricultural settings has prompted many hemipteran pests to develop resistance to these chemicals. This situation underscores the urgent need for the development of new pest control methods that are both species-specific and environmentally friendly. Notably, the field of hemipteran biotechnology has made significant progress, with CRISPR/Cas-mediated hemipteran mutagenesis techniques gaining prominence [130].

2.3.1. Bemisia tabaci

B. tabaci is a serious agricultural polyphagous pest and a carrier of several plant viruses that cause significant economic losses worldwide [131]. Traditional embryo injection methods are technically difficult and have high mortality rates, for example, the A2083V mutation associated with ketoenol resistance was detected using CRISPR/Cas9 introduced into D. melanogaster [112]. Thus, researchers have developed a new CRISPR/Cas9 gene-editing program based on the injection of vitellogenic adult females to form “ReMOT Control”. The protocol uses an ovary-targeting peptide ligand (BtKV) to fuse and inject Cas9 into adult females, enabling efficient, heritable editing of the offspring genome. This adult injection method is simple and easy to perform, requires no specialized equipment, and opens up new possibilities for gene-editing studies in B. tabaci [132].

2.3.2. Nilaparvata lugens

N. lugens serves as the primary pest of rice crops in China and various Asian nations, relying exclusively on rice and wild rice for feeding and reproduction [133]. The resistance molecular mechanisms of N. lugens to multiple insecticides included P450-mediated metabolic detoxification as a major pathway. CYP6ER1 overexpression is an important resistance mechanism of N. lugens to neonicotinoid insecticides. Zhang utilized CRISPR/Cas9 technology to create a CYP6ER1-knockout homozygous strain, revealing a significant increase in sensitivity to imidacloprid and thiacloprid, while the increase in sensitivity to other neonicotinoids was comparatively modest. Metabolic analyses and site predictions indicate that CYP6ER1 activity correlates with insecticide structure, showing that imidacloprid and thiacloprid are readily oxidized in a specific five-membered heterocycle [110]. Zhang knocked out CYP6CS1 to investigate its influence on resistance to various insecticides, finding a marked enhancement in sensitivity to imidacloprid and other insecticides. However, differences were noted in survival rates, reproduction, and weight between the CYP6CS1KO strain and the wild type [111]. A single non-synonymous SNP was found in the chitin synthase 1 gene (CHS1) at the amino acid position of 932, where a glycine (G) was replaced with cysteine (C), and scientists demonstrated that this single-amino acid replacement is associated with buprofezin resistance. At the same position, Zhang introduced the substitution mutations (G932C) of the CHS1 gene into N. lugens using the CRISPR/Cas9 system combined with the HDR pathway. The N. lugens lines that are homozygous for these mutations (G932C) were resistant to buprofezin [134].

2.3.3. Myzus persicae

Aphidoidea is an insect that is a serious hazard to agriculture and requires effective control measures to reduce its damage. Simultaneously, Aphidoidea also has some special biological phenomena and ecological relationships that deserve further research and exploration [135]. Resistance of M. persicae to neonicotinoid insecticides is mainly caused by an arginine-to-threonine substitution at position 81 of the nAChR_beta 1 subunit (R81T mutation). This mutation confers tolerance to neonicotinoid insecticides in M. persicae. To validate the role of R81T in neonicotinoid resistance and to test whether it imposes any significant fitness costs on the insect, Homem used CRISPR/Cas9 to introduce a similar mutation in the genome of D. melanogaster and found that D. melanogaster carrying R81T showed increased resistance to neonicotinoid insecticides accompanied by a significant reduction in fitness [113]. In a follow-up study, a novel RPA-CRISPR/Cas12a-based rapid visual assay was developed to specifically target the nAChR subunit V62I and R81T mutations in Aphis gossypii to monitor its resistance to imidacloprid. The method uses A. gossypii tissue samples of less than 2 mm to accurately identify imidacloprid-resistant A. gossypii populations in the field in less than 1 h at 37 °C and 440–460 nm blue light [127].

3. Conclusions and Prospects

As the global population continues to grow and the need for food becomes more urgent, agricultural pests have become a serious challenge affecting food security and agricultural production. These pests not only cause heavy crop losses worldwide, but also pose a threat to human health by transmitting bacteria and viruses [136,137]. To address this challenge, scientists have been seeking effective pest control strategies. However, conventional insecticides have led to the gradual development of pest resistance due to overuse, which greatly reduces the effectiveness of pest management. Therefore, it has become particularly urgent to gain a deeper understanding of the mechanisms of insect resistance and to develop new pest control methods. The emergence of CRISPR/Cas technology has provided new perspectives and powerful tools for entomological research. Especially in exploring insect resistance, CRISPR/Cas technology not only helps to reveal the molecular mechanisms of insect resistance, but also offers the possibility of developing new pest management strategies.
In insect drug resistance research, CRISPR/Cas technology is widely used in a variety of insect models, such as D. melanogaster. Through CRISPR/Cas9 technology, researchers are able to precisely edit the genome of D melanogaster to explore the genes and pathways associated with drug resistance. For example, CYP genes, which are key members of insect metabolic detoxification enzymes, were identified in D melanogaster as important candidate genes associated with multiple insecticide metabolism and resistance [51,52]. In addition, nAChR and ABC transporter proteins also play important roles in insect resistance, and the knockdown or mutation of these genes by CRISPR/Cas9 technology can further reveal their resistance mechanisms [56,59]. In addition to D melanogaster, CRISPR/Cas technology has also been applied in the study of drug resistance in other agricultural pests, such as H. armigera [85,90], S. exigua [99,102], and P. xylostella [65,99]. In these pests, researchers knocked out resistance-related genes, such as cadherin, P450 gene, and GST gene, by CRISPR/Cas9 technology, thus revealing the role of these genes in pest resistance. In addition, the resistance mutants of pests can be created by CRISPR/Cas technology, providing new tools and methods for monitoring and management of pest resistance. By editing these gene, scientists can identify genes associated with insect resistance and, through the intensive study of these genes, further reduce the number of pests and the damage they cause to crops. In the control of public health pests, CRISPR technology also has broad application prospects. For example, Ae. Aegypti is a mosquito that is a major vector of serious infectious diseases such as dengue fever. With CRISPR technology, scientists can edit the genes of the Ae. aegypti to make it unable to carry and transmit viruses, thus blocking the spread of disease at the source [61,119].
CRISPR holds a substantial promise for leveraging its capabilities in insecticidal interventions and innovative approaches to pest management. In recent years, a team from the University of California has established a platform for precision-guided sterile insect technique (pgSIT) based on CRISPR and achieved remarkable results. They successfully created sterile insects by programming CRISPR/Cas9 to induce specific gene mutation sites, which resulted in a male infertility rate of over 99.5% and a female mortality rate of over 99.9% in hybrid offspring. This technique has been tested on a variety of pests including D. melanogaster and An. gambiae, among others [138,139]. DIPA-CRISPR (Direct Injection of Parental Animals for CRISPR) represents an innovative gene-editing technology. It involves directly injecting CRISPR/Cas9 components into the bodies of adult female insects. This method influences developing oocytes, allowing heritable genetic mutations in offspring, thereby simplifying the gene-editing process in insects [140]. However, to date, testing has occurred only in a limited number of insect species. Future advancements promise to enhance the efficiency and convenience of gene editing [141,142,143,144]. For populations that have developed resistance, CRISPR/Cas9 presents a potential solution—reversing resistance. Through gene drive technology, researchers can propagate specific genetic variations within insect populations, restoring sensitivity to insecticides [145]. Beyond reversing resistance, this technology also holds promise for the development of novel insecticides. By editing genes associated with drug metabolism, drug targets, or drug transport within insects, researchers can devise efficient, low-toxicity chemical or biological insecticides tailored to specific insect species [146]. Although the application of the CRISPR/Cas9 system to insecticide resistance in insects holds great promise, there are obvious challenges. One of the foremost concerns is off-target effects, whereby the system may inadvertently modify genes similar to the intended target. Researchers are adopting several strategies to address this issue, such as developing high-precision, low-off-target Cas9 variants and optimizing gRNA design with bioinformatics tools to ensure precise targeting. Technological developments have resulted in fewer off-target modifications, enhancing the safety and efficacy of applications [147].
Understanding the mechanisms by which insects develop resistance to insecticides can effectively prevent and control pests, significantly improving crop yield and quality, reducing crop losses, and ultimately increasing economic benefits. It can also reduce the number of pests such as mosquitoes and flies, thereby reducing the risk of disease transmission and ensuring human health [148,149].

Author Contributions

Conceptualization, Q.X. and M.W.; methodology, Q.X. and X.S.; software, M.W.; validation, J.Z., H.S. and X.W.; formal analysis, H.J. and X.W.; investigation, D.S.; resources, M.W.; data curation, H.S.; writing—original draft preparation, H.J. and X.W.; writing—review and editing, Q.X. and H.S; visualization, J.Z.; supervision, D.S.; project administration, X.S.; funding acquisition, D.S. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the National Natural Science Foundation of China (32302354, U21A20223), Zhejiang Provincial Programs for Natural Science (LQ24C140002), the Zhejiang Provincial Programs for Science and Technology Development (2023SNJF034), and the “Pioneer” and “Leading Goose” R&D Program of Zhejiang (No. 2022C02047 and 2023C02030).

Data Availability Statement

Data are contained within the article.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Telleria, E.L.; Martins-da-Silva, A.; Tempone, A.J.; Traub-Csekö, Y.M. Leishmania, microbiota and sand fly immunity. Parasitology 2018, 145, 1336–1353. [Google Scholar] [CrossRef] [PubMed]
  2. Fiallo-Olivé, E.; Pan, L.L.; Liu, S.S.; Navas-Castillo, J. Transmission of Begomoviruses and Other Whitefly-Borne Viruses: Dependence on the Vector Species. Phytopathology 2020, 110, 10–17. [Google Scholar] [CrossRef] [PubMed]
  3. Li, M.; Zhou, Y.; Cheng, J.; Wang, Y.; Lan, C.; Shen, Y. Response of the mosquito immune system and symbiotic bacteria to pathogen infection. Parasites Vectors 2024, 17, 69. [Google Scholar] [CrossRef]
  4. Norris, E.J.; Coats, J.R. Current and Future Repellent Technologies: The Potential of Spatial Repellents and Their Place in Mosquito-Borne Disease Control. Int. J. Environ. Res. Public Health 2017, 14, 124. [Google Scholar] [CrossRef]
  5. Alyokhin, A.; Chen, Y. Adaptation to toxic hosts as a factor in the evolution of insecticide resistance. Curr. Opin. Insect Sci. 2017, 21, 33–38. [Google Scholar] [CrossRef] [PubMed]
  6. Moyes, C.L.; Vontas, J.; Martins, A.J.; Ng, L.C.; Koou, S.Y.; Dusfour, I.; Raghavendra, K.; Pinto, J.; Corbel, V.; David, J.P.; et al. Contemporary status of insecticide resistance in the major Aedes vectors of arboviruses infecting humans. PLoS Neglected Trop. Dis. 2017, 11, e0005625. [Google Scholar] [CrossRef]
  7. Jayapal, P.K.; Joshi, R.; Sathasivam, R.; Nguyen, B.V.; Faqeerzada, M.A.; Park, S.U.; Sandanam, D.; Cho, B.K. Non-destructive measurement of total phenolic compounds in Arabidopsis under various stress conditions. Front. Plant Sci. 2022, 13, 982247. [Google Scholar] [CrossRef]
  8. Zhang, B.; Li, C.; Luan, Y.; Lu, Y.; Hu, H.; Liu, Y.; Lu, K.; Zhang, G.; Dai, F.; Tong, X. The Role of Chitooligosaccharidolytic β-N-Acetylglucosamindase in the Molting and Wing Development of the Silkworm Bombyx mori. Int. J. Mol. Sci. 2022, 23, 3850. [Google Scholar] [CrossRef]
  9. Burc, E.; Girard-Tercieux, C.; Metz, M.; Cazaux, E.; Baur, J.; Koppik, M.; Rêgo, A.; Hart, A.F.; Berger, D. Life-history adaptation under climate warming magnifies the agricultural footprint of a cosmopolitan insect pest. Nat. Commun. 2025, 16, 827. [Google Scholar] [CrossRef]
  10. Nie, D.; Li, J.; Xie, Q.; Ai, L.; Zhu, C.; Wu, Y.; Gui, Q.; Zhang, L.; Tan, W. Nanoparticles: A Potential and Effective Method to Control Insect-Borne Diseases. Bioinorg. Chem. Appl. 2023, 2023, 5898160. [Google Scholar] [CrossRef]
  11. Araujo, S.; Mabille, D.; Garcia, A.B.; Caljon, G. A breath of fresh air: Impact of insect-borne protozoan parasites on the respiratory system. Trends Parasitol. 2024, 40, 717–730. [Google Scholar] [CrossRef]
  12. Al-Osaimi, H.M.; Kanan, M.; Marghlani, L.; Al-Rowaili, B.; Albalawi, R.; Saad, A.; Alasmari, S.; Althobaiti, K.; Alhulaili, Z.; Alanzi, A.; et al. A systematic review on malaria and dengue vaccines for the effective management of these mosquito borne diseases: Improving public health. Hum. Vaccines Immunother. 2024, 20, 2337985. [Google Scholar] [CrossRef] [PubMed]
  13. Tang, B.; Xu, K.; Liu, Y.; Zhou, Z.; Karthi, S.; Yang, H.; Li, C. A review of physiological resistance to insecticide stress in Nilaparvata lugens. 3 Biotech 2022, 12, 84. [Google Scholar] [CrossRef] [PubMed]
  14. Gnankiné, O.; Bassolé, I.H.; Chandre, F.; Glitho, I.; Akogbeto, M.; Dabiré, R.K.; Martin, T. Insecticide resistance in Bemisia tabaci Gennadius (Homoptera: Aleyrodidae) and Anopheles gambiae Giles (Diptera: Culicidae) could compromise the sustainability of malaria vector control strategies in West Africa. Acta Trop. 2013, 128, 7–17. [Google Scholar] [CrossRef] [PubMed]
  15. Oliva, C.F.; Vreysen, M.J.; Dupé, S.; Lees, R.S.; Gilles, J.R.; Gouagna, L.C.; Chhem, R. Current status and future challenges for controlling malaria with the sterile insect technique: Technical and social perspectives. Acta Trop. 2014, 132, S130–S139. [Google Scholar] [CrossRef] [PubMed]
  16. Gott, R.C.; Kunkel, G.R.; Zobel, E.S.; Lovett, B.R.; Hawthorne, D.J. Implicating ABC Transporters in Insecticide Resistance: Research Strategies and a Decision Framework. J. Econ. Entomol. 2017, 110, 667–677. [Google Scholar] [CrossRef]
  17. Reid, M.C.; McKenzie, F.E. The contribution of agricultural insecticide use to increasing insecticide resistance in African malaria vectors. Malar. J. 2016, 15, 107. [Google Scholar] [CrossRef]
  18. Zuo, Y.; Shi, Y.; Zhang, F.; Guan, F.; Zhang, J.; Feyereisen, R.; Fabrick, J.A.; Yang, Y.; Wu, Y. Genome mapping coupled with CRISPR gene editing reveals a P450 gene confers avermectin resistance in the beet armyworm. PLoS Genet. 2021, 17, e1009680. [Google Scholar] [CrossRef]
  19. Hu, B.; Huang, H.; Hu, S.; Ren, M.; Wei, Q.; Tian, X.; Esmail Abdalla Elzaki, M.; Bass, C.; Su, J.; Reddy Palli, S. Changes in both trans- and cis-regulatory elements mediate insecticide resistance in a lepidopteron pest, Spodoptera exigua. PLoS Genet. 2021, 17, e1009403. [Google Scholar] [CrossRef]
  20. Kaplanoglu, E.; Chapman, P.; Scott, I.M.; Donly, C. Overexpression of a cytochrome P450 and a UDP-glycosyltransferase is associated with imidacloprid resistance in the Colorado potato beetle, Leptinotarsa decemlineata. Sci. Rep. 2017, 7, 10. [Google Scholar] [CrossRef]
  21. Qin, J.; Yuchi, Z. Identification of a Novel Inhibitor of Cimex lectularius Acetylcholinesterase. J. Agric. Food Chem. 2024, 72, 12498–12507. [Google Scholar] [CrossRef] [PubMed]
  22. Roberts, P.H.; Zhou, X.; Holmes, A.M.; Ranson, H.; Small, G.; Hemingway, J.; Ng, J.D.; Chen, L.; Meehan, E.J. Crystallization of agGST1-6, a recombinant glutathione S-transferase from a DDT-resistant strain of Anopheles gambiae. Acta Crystallogr. D Biol. Crystallogr. 2001, 57, 134–136. [Google Scholar] [CrossRef] [PubMed]
  23. Gong, M.; Gu, Y.; Hu, X.; Sun, Y.; Ma, L.; Li, X.; Sun, L.; Sun, J.; Qian, J.; Zhu, C. Cloning and overexpression of CYP6F1, a cytochrome P450 gene, from deltamethrin-resistant Culex pipiens pallens. Acta Biochim. Biophys. Sin. 2005, 37, 317–326. [Google Scholar] [CrossRef] [PubMed]
  24. Ren, X.; Han, Z.; Wang, Y. Mechanisms of monocrotophos resistance in cotton bollworm, Helicoverpa armigera (Hübner). Arch. Insect Biochem. Physiol. 2002, 51, 103–110. [Google Scholar] [CrossRef] [PubMed]
  25. Muthusamy, R.; Shivakumar, M.S. Involvement of metabolic resistance and F1534C kdr mutation in the pyrethroid resistance mechanisms of Aedes aegypti in India. Acta Trop. 2015, 148, 137–141. [Google Scholar] [CrossRef]
  26. Sun, D.; Guo, Z.; Liu, Y.; Zhang, Y. Progress and Prospects of CRISPR/Cas Systems in Insects and Other Arthropods. Front. Physiol. 2017, 8, 608. [Google Scholar] [CrossRef]
  27. Riordan, S.M.; Heruth, D.P.; Zhang, L.; Ye, S. Application of CRISPR/Cas9 for biomedical discoveries. Cell Biosci. 2015, 5, 33. [Google Scholar] [CrossRef]
  28. Merlin, C.; Beaver, L.E.; Taylor, O.R.; Wolfe, S.A.; Reppert, S.M. Efficient targeted mutagenesis in the monarch butterfly using zinc-finger nucleases. Genome Res. 2013, 23, 159–168. [Google Scholar] [CrossRef]
  29. Jo, Y.I.; Kim, H.; Ramakrishna, S. Recent developments and clinical studies utilizing engineered zinc finger nuclease technology. Cell. Mol. Life Sci. 2015, 72, 3819–3830. [Google Scholar] [CrossRef]
  30. Bibikova, M.; Golic, M.; Golic, K.G.; Carroll, D. Targeted chromosomal cleavage and mutagenesis in Drosophila using zinc-finger nucleases. Genetics 2002, 161, 1169–1175. [Google Scholar] [CrossRef]
  31. Beumer, K.; Bhattacharyya, G.; Bibikova, M.; Trautman, J.K.; Carroll, D. Efficient gene targeting in Drosophila with zinc-finger nucleases. Genetics 2006, 172, 2391–2403. [Google Scholar] [CrossRef] [PubMed]
  32. Carroll, D.; Beumer, K.J.; Morton, J.J.; Bozas, A.; Trautman, J.K. Gene targeting in Drosophila and Caenorhabditis elegans with zinc-finger nucleases. Methods Mol. Biol. 2008, 435, 63–77. [Google Scholar] [PubMed]
  33. Sajwan, S.; Takasu, Y.; Tamura, T.; Uchino, K.; Sezutsu, H.; Zurovec, M. Efficient disruption of endogenous Bombyx gene by TAL effector nucleases. Insect Biochem. Mol. Biol. 2013, 43, 17–23. [Google Scholar] [CrossRef]
  34. Mock, U.; Machowicz, R.; Hauber, I.; Horn, S.; Abramowski, P.; Berdien, B.; Hauber, J.; Fehse, B. mRNA transfection of a novel TAL effector nuclease (TALEN) facilitates efficient knockout of HIV co-receptor CCR5. Nucleic Acids Res. 2015, 43, 5560–5571. [Google Scholar] [CrossRef] [PubMed]
  35. Liu, J.; Li, C.; Yu, Z.; Huang, P.; Wu, H.; Wei, C.; Zhu, N.; Shen, Y.; Chen, Y.; Zhang, B.; et al. Efficient and specific modifications of the Drosophila genome by means of an easy TALEN strategy. J. Genet. Genom. 2012, 39, 209–215. [Google Scholar] [CrossRef] [PubMed]
  36. Richter, H.; Randau, L.; Plagens, A. Exploiting CRISPR/Cas: Interference mechanisms and applications. Int. J. Mol. Sci. 2013, 14, 14518–14531. [Google Scholar] [CrossRef]
  37. 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]
  38. Lander, E.S. The Heroes of CRISPR. Cell 2016, 164, 18–28. [Google Scholar] [CrossRef]
  39. Lai, S.; Wei, S.; Zhao, B.; Ouyang, Z.; Zhang, Q.; Fan, N.; Liu, Z.; Zhao, Y.; Yan, Q.; Zhou, X.; et al. Generation of Knock-In Pigs Carrying Oct4-tdTomato Reporter through CRISPR/Cas9-Mediated Genome Engineering. PLoS ONE 2016, 11, e0146562. [Google Scholar] [CrossRef]
  40. Lamb, A.M.; Walker, E.A.; Wittkopp, P.J. Tools and strategies for scarless allele replacement in Drosophila using CRISPR/Cas9. Fly 2017, 11, 53–64. [Google Scholar] [CrossRef]
  41. Zhu, S.; Yu, X.; Li, Y.; Sun, Y.; Zhu, Q.; Sun, J. Highly Efficient Targeted Gene Editing in Upland Cotton Using the CRISPR/Cas9 System. Int. J. Mol. Sci. 2018, 19, 3000. [Google Scholar] [CrossRef] [PubMed]
  42. Yan, Y.; Aumann, R.A.; Häcker, I.; Schetelig, M.F. CRISPR-based genetic control strategies for insect pests. J. Integr. Agric. 2023, 22, 651–668. [Google Scholar] [CrossRef]
  43. Itokawa, K.; Komagata, O.; Kasai, S.; Ogawa, K.; Tomita, T. Testing the causality between CYP9M10 and pyrethroid resistance using the TALEN and CRISPR/Cas9 technologies. Sci. Rep. 2016, 6, 24652. [Google Scholar] [CrossRef] [PubMed]
  44. Scott, J.G.; Michel, K.; Bartholomay, L.C.; Siegfried, B.D.; Hunter, W.B.; Smagghe, G.; Zhu, K.Y.; Douglas, A.E. Towards the elements of successful insect RNAi. J. Insect Physiol. 2013, 59, 1212–1221. [Google Scholar] [CrossRef]
  45. Perkin, L.C.; Adrianos, S.L.; Oppert, B. Gene Disruption Technologies Have the Potential to Transform Stored Product Insect Pest Control. Insects 2016, 7, 46. [Google Scholar] [CrossRef] [PubMed]
  46. Gratz, S.J.; Cummings, A.M.; Nguyen, J.N.; Hamm, D.C.; Donohue, L.K.; Harrison, M.M.; Wildonger, J.; O’Connor-Giles, K.M. Genome engineering of Drosophila with the CRISPR RNA-guided Cas9 nuclease. Genetics 2013, 194, 1029–1035. [Google Scholar] [CrossRef] [PubMed]
  47. 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]
  48. Yang, J.; Schleicher, T.R.; Dong, Y.; Park, H.B.; Lan, J.; Cresswell, P.; Crawford, J.; Dimopoulos, G.; Fikrig, E. Disruption of mosGILT in Anopheles gambiae impairs ovarian development and Plasmodium infection. J. Exp. Med. 2020, 217, e20190682. [Google Scholar] [CrossRef]
  49. Nateghi Rostami, M. CRISPR/Cas9 gene drive technology to control transmission of vector-borne parasitic infections. Parasite Immunol. 2020, 42, e12762. [Google Scholar] [CrossRef]
  50. Yu, Y.; Liu, X.J.; Ma, X.; Zhang, Z.J.; Wang, T.C.; Sun, F.; Hou, C.X.; Li, M.W. A palmitoyltransferase Approximated gene Bm-app regulates wing development in Bombyx mori. Insect Sci. 2020, 27, 2–13. [Google Scholar] [CrossRef]
  51. Denecke, S.; Fusetto, R.; Martelli, F.; Giang, A.; Battlay, P.; Fournier-Level, A.; RA, O.H.; Batterham, P. Multiple P450s and Variation in Neuronal Genes Underpins the Response to the Insecticide Imidacloprid in a Population of Drosophila melanogaster. Sci. Rep. 2017, 7, 11338. [Google Scholar] [CrossRef] [PubMed]
  52. Fusetto, R.; Denecke, S.; Perry, T.; O’Hair, R.A.J.; Batterham, P. Partitioning the roles of CYP6G1 and gut microbes in the metabolism of the insecticide imidacloprid in Drosophila melanogaster. Sci. Rep. 2017, 7, 11339. [Google Scholar] [CrossRef] [PubMed]
  53. Kaduskar, B.; Kushwah, R.B.; Auradkar, A.; Guichard, A.; Li, M.; Bennett, J.B.; Julio, A.H.; Marshall, J.M.; Montell, C.; Bier, E. Reversing insecticide resistance with allelic-drive in Drosophila melanogaster. Nat. Commun. 2022, 13, 291. [Google Scholar] [CrossRef]
  54. Samantsidis, G.R.; Panteleri, R.; Denecke, S.; Kounadi, S.; Christou, I.; Nauen, R.; Douris, V.; Vontas, J. ‘What I cannot create, I do not understand’: Functionally validated synergism of metabolic and target site insecticide resistance. Proc. Biol. Sci. 2020, 287, 20200838. [Google Scholar] [CrossRef]
  55. Zimmer, C.T.; Garrood, W.T.; Puinean, A.M.; Eckel-Zimmer, M.; Williamson, M.S.; Davies, T.G.; Bass, C. A CRISPR/Cas9 mediated point mutation in the alpha 6 subunit of the nicotinic acetylcholine receptor confers resistance to spinosad in Drosophila melanogaster. Insect Biochem. Mol. Biol. 2016, 73, 62–69. [Google Scholar] [CrossRef] [PubMed]
  56. Perry, T.; Chen, W.; Ghazali, R.; Yang, Y.T.; Christesen, D.; Martelli, F.; Lumb, C.; Bao Luong, H.N.; Mitchell, J.; Holien, J.K.; et al. Role of nicotinic acetylcholine receptor subunits in the mode of action of neonicotinoid, sulfoximine and spinosyn insecticides in Drosophila melanogaster. Insect Biochem. Mol. Biol. 2021, 131, 103547. [Google Scholar] [CrossRef]
  57. Hernandes, N.; Qi, X.; Bhide, S.; Brown, C.; Camm, B.J.; Baxter, S.W.; Robin, C. Acetylcholine esterase of Drosophila melanogaster: A laboratory model to explore insecticide susceptibility gene drives. Pest Manag. Sci. 2024, 80, 2950–2964. [Google Scholar] [CrossRef]
  58. Vernon, S.W.; Goodchild, J.; Baines, R.A. The VAChTY49N mutation provides insecticide-resistance but perturbs evoked cholinergic neurotransmission in Drosophila. PLoS ONE 2018, 13, e0203852. [Google Scholar] [CrossRef] [PubMed]
  59. Denecke, S.; Fusetto, R.; Batterham, P. Describing the role of Drosophila melanogaster ABC transporters in insecticide biology using CRISPR-Cas9 knockouts. Insect Biochem. Mol. Biol. 2017, 91, 1–9. [Google Scholar] [CrossRef]
  60. Zhou, T.; Wu, W.; Ma, S.; Chen, J.; Huang, J.; Qiao, X. Effects of RDL GABA Receptor Point Mutants on Susceptibility to Meta-Diamide and Isoxazoline Insecticides in Drosophila melanogaster. Insects 2024, 15, 334. [Google Scholar] [CrossRef]
  61. Pacheco, S.; Gallegos, A.S.; Peláez-Aguilar, Á.E.; Sánchez, J.; Gómez, I.; Soberón, M.; Bravo, A. CRISPR-Cas9 knockout of membrane-bound alkaline phosphatase or cadherin does not confer resistance to Cry toxins in Aedes aegypti. PLoS Neglected Trop. Dis. 2024, 18, e0012256. [Google Scholar] [CrossRef] [PubMed]
  62. Feng, X.; López Del Amo, V.; Mameli, E.; Lee, M.; Bishop, A.L.; Perrimon, N.; Gantz, V.M. Optimized CRISPR tools and site-directed transgenesis towards gene drive development in Culex quinquefasciatus mosquitoes. Nat. Commun. 2021, 12, 2960. [Google Scholar] [CrossRef] [PubMed]
  63. Grigoraki, L.; Cowlishaw, R.; Nolan, T.; Donnelly, M.; Lycett, G.; Ranson, H. CRISPR/Cas9 modified An. gambiae carrying kdr mutation L1014F functionally validate its contribution in insecticide resistance and combined effect with metabolic enzymes. PLoS Genet. 2021, 17, e1009556. [Google Scholar] [CrossRef]
  64. Williams, J.; Cowlishaw, R.; Sanou, A.; Ranson, H.; Grigoraki, L. In vivo functional validation of the V402L voltage gated sodium channel mutation in the malaria vector An. gambiae. Pest Manag. Sci. 2022, 78, 1155–1163. [Google Scholar] [CrossRef] [PubMed]
  65. 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]
  66. Guo, Z.J.; Guo, L.; Qin, J.Y.; Ye, F.; Sun, D.; Wu, Q.J.; Wang, S.L.; Crickmore, N.; Zhou, X.G.; Bravo, A.; et al. A single transcription factor facilitates an insect host combating Bacillus thuringiensis infection while maintaining fitness. Nat. Commun. 2022, 13, 15. [Google Scholar] [CrossRef] [PubMed]
  67. Guo, Z.J.; Kang, S.; Sun, D.; Gong, L.J.; Zhou, J.L.; Qin, J.Y.; Guo, L.; Zhu, L.H.; Bai, Y.; Ye, F.; et al. MAPK-dependent hormonal signaling plasticity contributes to overcoming Bacillus thuringiensis toxin action in an insect host. Nat. Commun. 2020, 11, 14. [Google Scholar] [CrossRef]
  68. 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]
  69. Sun, D.; Xu, Q.; Guo, L.; Bai, Y.; Shentu, X.; Yu, X.; Crickmore, N.; Zhou, X.; Bravo, A.; Soberón, M.; et al. The role of GPI-anchored membrane-bound alkaline phosphatase in the mode of action of Bt Cry1A toxins in the diamondback moth. Fundam. Res. 2024; In Press, Corrected Proof. [Google Scholar] [CrossRef]
  70. Liu, Z.; Fu, S.; Ma, X.; Baxter, S.W.; Vasseur, L.; Xiong, L.; Huang, Y.; Yang, G.; You, S.; You, M. Resistance to Bacillus thuringiensis Cry1Ac toxin requires mutations in two Plutella xylostella ATP-binding cassette transporter paralogs. PLoS Pathog. 2020, 16, e1008697. [Google Scholar] [CrossRef]
  71. 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]
  72. Ouyang, C.Z.; Ye, F.; Wu, Q.J.; Wang, S.L.; Crickmore, N.; Zhou, X.G.; Guo, Z.J.; Zhang, Y.J. CRISPR/Cas9-based functional characterization of PxABCB1 reveals its roles in the resistance of Plutella xylostella (L.) to Cry1Ac, abamectin and emamectin benzoate. J. Integr. Agric. 2023, 22, 3090–3102. [Google Scholar] [CrossRef]
  73. 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]
  74. 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]
  75. Guo, Z.J.; Guo, L.; Bai, Y.; Kang, S.; Sun, D.; Qin, J.Y.; Ye, F.; Wang, S.L.; Wu, Q.J.; Xie, W.; et al. Retrotransposon-mediated evolutionary rewiring of a pathogen response orchestrates a resistance phenotype in an insect host. Proc. Natl. Acad. Sci. USA 2023, 120, 10. [Google Scholar] [CrossRef] [PubMed]
  76. Guo, Z.; Zhu, L.; Cheng, Z.; Dong, L.; Guo, L.; Bai, Y.; Wu, Q.; Wang, S.; Yang, X.; Xie, W.; et al. A midgut transcriptional regulatory loop favors an insect host to withstand a bacterial pathogen. Innovation 2024, 5, 100675. [Google Scholar] [CrossRef] [PubMed]
  77. Douris, V.; Steinbach, D.; Panteleri, R.; Livadaras, I.; Pickett, J.A.; Van Leeuwen, T.; Nauen, R.; Vontas, J. Resistance mutation conserved between insects and mites unravels the benzoylurea insecticide mode of action on chitin biosynthesis. Proc. Natl. Acad. Sci. USA 2016, 113, 14692–14697. [Google Scholar] [CrossRef]
  78. Guo, Z.J.; Bai, Y.; Zhang, X.Y.; Guo, L.; Zhu, L.H.; Sun, D.; Sun, K.Y.; Xu, X.D.; Yang, X.; Xie, W.; et al. RNA m6A Methylation Suppresses Insect Juvenile Hormone Degradation to Minimize Fitness Costs in Response to A Pathogenic Attack. Adv. Sci. 2024, 11, 17. [Google Scholar] [CrossRef]
  79. 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. 2020, 76, 1618–1625. [Google Scholar] [CrossRef]
  80. 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]
  81. Sun, X.; Hua, W.; Zhu, B.; Liang, P.; Gao, X. CRISPR/Cas9-mediated D472N substitution in the Rdl1 of Plutella xylostella confers low resistance to abamectin. Pest Manag. Sci. 2023, 79, 741–747. [Google Scholar] [CrossRef] [PubMed]
  82. 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] [PubMed]
  83. Jiang, D.; Yu, Z.; He, Y.; Wang, F.; Gu, Y.; Davies, T.G.; 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]
  84. Huang, J.M.; Rao, C.; Wang, S.; He, L.F.; Zhao, S.Q.; Zhou, L.Q.; Zhao, Y.X.; Yang, F.X.; Gao, C.F.; Wu, S.F. Multiple target-site mutations occurring in lepidopterans confer resistance to diamide insecticides. Insect Biochem. Mol. Biol. 2020, 121, 103367. [Google Scholar] [CrossRef] [PubMed]
  85. Wang, J.; Zhang, H.; Wang, H.; Zhao, S.; Zuo, Y.; Yang, Y.; Wu, Y. Functional validation of cadherin as a receptor of Bt toxin Cry1Ac in Helicoverpa armigera utilizing the CRISPR/Cas9 system. Insect Biochem. Mol. Biol. 2016, 76, 11–17. [Google Scholar] [CrossRef]
  86. Wang, J.; Zhao, X.; Yan, R.; Wu, S.; Wu, Y.; Yang, Y. Reverse genetics reveals contrary effects of two Rdl-homologous GABA receptors of Helicoverpa armigera on the toxicity of cyclodiene insecticides. Pestic. Biochem. Physiol. 2020, 170, 104699. [Google Scholar] [CrossRef]
  87. Wang, J.; Ma, H.; Zuo, Y.; Yang, Y.; Wu, Y. CRISPR-mediated gene knockout reveals nicotinic acetylcholine receptor (nAChR) subunit α6 as a target of spinosyns in Helicoverpa armigera. Pest Manag. Sci. 2020, 76, 2925–2931. [Google Scholar] [CrossRef]
  88. Wang, J.; Wang, H.; Liu, S.; Liu, L.; Tay, W.T.; Walsh, T.K.; Yang, Y.; Wu, Y. CRISPR/Cas9 mediated genome editing of Helicoverpa armigera with mutations of an ABC transporter gene HaABCA2 confers resistance to Bacillus thuringiensis Cry2A toxins. Insect Biochem. Mol. Biol. 2017, 87, 147–153. [Google Scholar] [CrossRef]
  89. Zhang, D.; Jin, M.; Yang, Y.; Zhang, J.; Yang, Y.; Liu, K.; Soberón, M.; Bravo, A.; Xiao, Y.; Wu, K. Synergistic resistance of Helicoverpa armigera to Bt toxins linked to cadherin and ABC transporters mutations. Insect Biochem. Mol. Biol. 2021, 137, 103635. [Google Scholar] [CrossRef]
  90. Jin, M.; Peng, Y.; Peng, J.; Zhang, H.; Shan, Y.; Liu, K.; Xiao, Y. Transcriptional regulation and overexpression of GST cluster enhances pesticide resistance in the cotton bollworm, Helicoverpa armigera (Lepidoptera: Noctuidae). Commun. Biol. 2023, 6, 1064. [Google Scholar] [CrossRef]
  91. Fabrick, J.A.; LeRoy, D.M.; Mathew, L.G.; Wu, Y.; Unnithan, G.C.; Yelich, A.J.; Carrière, Y.; Li, X.; Tabashnik, B.E. CRISPR-mediated mutations in the ABC transporter gene ABCA2 confer pink bollworm resistance to Bt toxin Cry2Ab. Sci. Rep. 2021, 11, 10377. [Google Scholar] [CrossRef] [PubMed]
  92. Fabrick, J.A.; Heu, C.C.; LeRoy, D.M.; DeGain, B.A.; Yelich, A.J.; Unnithan, G.C.; Wu, Y.; Li, X.; Carrière, Y.; Tabashnik, B.E. Knockout of ABC transporter gene ABCA2 confers resistance to Bt toxin Cry2Ab in Helicoverpa zea. Sci. Rep. 2022, 12, 16706. [Google Scholar] [CrossRef] [PubMed]
  93. Xu, L.; Li, B.; Liu, H.; Zhang, H.; Liu, R.; Yu, H.; Li, D. CRISPR/Cas9-Mediated Knockout Reveals the Involvement of CYP304F1 in β-Cypermethrin and Chlorpyrifos Resistance in Spodoptera litura. J. Agric. Food Chem. 2022, 70, 11192–11200. [Google Scholar] [CrossRef] [PubMed]
  94. Liu, Z.; Liao, C.; Zou, L.; Jin, M.; Shan, Y.; Quan, Y.; Yao, H.; Zhang, L.; Wang, P.; Liu, Z.; et al. Retrotransposon-mediated disruption of a chitin synthase gene confers insect resistance to Bacillus thuringiensis Vip3Aa toxin. PLoS Biol. 2024, 22, e3002704. [Google Scholar] [CrossRef] [PubMed]
  95. Shi, T.; Tang, P.; Wang, X.; Yang, Y.; Wu, Y. CRISPR-mediated knockout of nicotinic acetylcholine receptor (nAChR) α6 subunit confers high levels of resistance to spinosyns in Spodoptera frugiperda. Pestic. Biochem. Physiol. 2022, 187, 105191. [Google Scholar] [CrossRef]
  96. Shi, Y.; Liu, Q.; Lu, W.; Yuan, J.; Yang, Y.; Oakeshott, J.; Wu, Y. Divergent amplifications of CYP9A cytochrome P450 genes provide two noctuid pests with differential protection against xenobiotics. Proc. Natl. Acad. Sci. USA 2023, 120, e2308685120. [Google Scholar] [CrossRef]
  97. Yang, X.; Chen, W.; Song, X.; Ma, X.; Cotto-Rivera, R.O.; Kain, W.; Chu, H.; Chen, Y.R.; Fei, Z.; Wang, P. Mutation of ABC transporter ABCA2 confers resistance to Bt toxin Cry2Ab in Trichoplusia ni. Insect Biochem. Mol. Biol. 2019, 112, 103209. [Google Scholar] [CrossRef]
  98. Ma, X.; Shao, E.; Chen, W.; Cotto-Rivera, R.O.; Yang, X.; Kain, W.; Fei, Z.; Wang, P. Bt Cry1Ac resistance in Trichoplusia ni is conferred by multi-gene mutations. Insect Biochem. Mol. Biol. 2022, 140, 103678. [Google Scholar] [CrossRef]
  99. Zuo, Y.; Wang, H.; Xu, Y.; Huang, J.; Wu, S.; Wu, Y.; Yang, Y. CRISPR/Cas9 mediated G4946E substitution in the ryanodine receptor of Spodoptera exigua confers high levels of resistance to diamide insecticides. Insect Biochem. Mol. Biol. 2017, 89, 79–85. [Google Scholar] [CrossRef]
  100. Zuo, Y.; Huang, J.; Wang, J.; Feng, Y.; Han, T.; Wu, Y.; Yang, Y. Knockout of a P-glycoprotein gene increases susceptibility to abamectin and emamectin benzoate in Spodoptera exigua. Insect Mol. Biol. 2018, 27, 36–45. [Google Scholar] [CrossRef]
  101. Zuo, Y.; Xue, Y.; Lu, W.; Ma, H.; Chen, M.; Wu, Y.; Yang, Y.; Hu, Z. Functional validation of nicotinic acetylcholine receptor (nAChR) α6 as a target of spinosyns in Spodoptera exigua utilizing the CRISPR/Cas9 system. Pest Manag. Sci. 2020, 76, 2415–2422. [Google Scholar] [CrossRef]
  102. Zuo, Y.; Xue, Y.; Wang, Z.; Ren, X.; Aioub, A.A.; Wu, Y.; Yang, Y.; Hu, Z. Knockin of the G275E mutation of the nicotinic acetylcholine receptor (nAChR) α6 confers high levels of resistance to spinosyns in Spodoptera exigua. Insect Sci. 2022, 29, 478–486. [Google Scholar] [CrossRef] [PubMed]
  103. Wang, Z.; Zhang, R.; Pei, Y.; Wu, W.; Hu, Z.; Zuo, Y. The knockout of the nicotinic acetylcholine receptor subunit gene α1 (nAChR α1) through CRISPR/Cas9 technology exposes its involvement in the resistance of Spodoptera exigua to insecticides. Pestic. Biochem. Physiol. 2023, 196, 105616. [Google Scholar] [CrossRef] [PubMed]
  104. Wang, X.; Xu, Y.; Huang, J.; Jin, W.; Yang, Y.; Wu, Y. CRISPR-Mediated Knockout of the ABCC2 Gene in Ostrinia furnacalis Confers High-Level Resistance to the Bacillus thuringiensis Cry1Fa Toxin. Toxins 2020, 12, 246. [Google Scholar] [CrossRef] [PubMed]
  105. Gao, Q.; Lin, Y.; Wang, X.; Jing, D.; Wang, Z.; He, K.; Bai, S.; Zhang, Y.; Zhang, T. Knockout of ABC Transporter ABCG4 Gene Confers Resistance to Cry1 Proteins in Ostrinia furnacalis. Toxins 2022, 14, 52. [Google Scholar] [CrossRef] [PubMed]
  106. Huang, J.; Sun, H.; He, L.; Liu, C.; Ge, W.; Ni, H.; Gao, C.; Wu, S. Double ryanodine receptor mutations confer higher diamide resistance in rice stem borer, Chilo suppressalis. Pest Manag. Sci. 2021, 77, 4971–4979. [Google Scholar] [CrossRef]
  107. Sun, H.; Wang, S.; Liu, C.; Hu, W.; Liu, J.; Zheng, L.; Gao, M.; Guo, F.; Qiao, S.; Liu, J.; et al. Risk assessment, fitness cost, cross-resistance, and mechanism of tetraniliprole resistance in the rice stem borer, Chilo suppressalis. Insect Sci. 2024, 31, 835–846. [Google Scholar] [CrossRef]
  108. Samantsidis, G.R.; O’Reilly, A.O.; Douris, V.; Vontas, J. Functional validation of target-site resistance mutations against sodium channel blocker insecticides (SCBIs) via molecular modeling and genome engineering in Drosophila. Insect Biochem. Mol. Biol. 2019, 104, 73–81. [Google Scholar] [CrossRef]
  109. Douris, V.; Papapostolou, K.M.; Ilias, A.; Roditakis, E.; Kounadi, S.; Riga, M.; Nauen, R.; Vontas, J. Investigation of the contribution of RyR target-site mutations in diamide resistance by CRISPR/Cas9 genome modification in Drosophila. Insect Biochem. Mol. Biol. 2017, 87, 127–135. [Google Scholar] [CrossRef]
  110. Zhang, H.; Zou, J.; Yang, B.; Zhang, Y.; Liu, Z. Importance of CYP6ER1 Was Different among Neonicotinoids in Their Susceptibility in Nilaparvata lugens. J. Agric. Food Chem. 2023, 71, 4163–4171. [Google Scholar] [CrossRef]
  111. Zhang, Y.; Gao, Y.; Ye, W.; Peng, Y.; Zhu, K.; Gao, C. CRISPR/Cas9-mediated knockout of NlCYP6CS1 gene reveals its role in detoxification of insecticides in Nilaparvata lugens (Hemiptera: Delphacidae). Pest Manag. Sci. 2023, 79, 2239–2246. [Google Scholar] [CrossRef]
  112. Lueke, B.; Douris, V.; Hopkinson, J.E.; Maiwald, F.; Hertlein, G.; Papapostolou, K.M.; Bielza, P.; Tsagkarakou, A.; Van Leeuwen, T.; Bass, C.; et al. Identification and functional characterization of a novel acetyl-CoA carboxylase mutation associated with ketoenol resistance in Bemisia tabaci. Pestic. Biochem. Physiol. 2020, 166, 104583. [Google Scholar] [CrossRef] [PubMed]
  113. Homem, R.A.; Buttery, B.; Richardson, E.; Tan, Y.; Field, L.M.; Williamson, M.S.; Davies, T.G.E. Evolutionary trade-offs of insecticide resistance—The fitness costs associated with target-site mutations in the nAChR of Drosophila melanogaster. Mol. Ecol. 2020, 29, 2661–2675. [Google Scholar] [CrossRef]
  114. Bassett, A.R.; Liu, J. CRISPR/Cas9 and genome editing in Drosophila. J. Genet. Genom. 2014, 41, 7–19. [Google Scholar] [CrossRef] [PubMed]
  115. Gao, H.; Cui, C.; Wang, L.; Jacobs-Lorena, M.; Wang, S. Mosquito Microbiota and Implications for Disease Control. Trends Parasitol. 2020, 36, 98–111. [Google Scholar] [CrossRef] [PubMed]
  116. Campos, E.V.; de Oliveira, J.L.; Abrantes, D.C.; Rogério, C.B.; Bueno, C.; Miranda, V.R.; Monteiro, R.A.; Fraceto, L.F. Recent Developments in Nanotechnology for Detection and Control of Aedes aegypti-Borne Diseases. Front. Bioeng. Biotechnol. 2020, 8, 102. [Google Scholar] [CrossRef] [PubMed]
  117. Amaral, J.K.; Bilsborrow, J.B.; Schoen, R.T. Chronic Chikungunya Arthritis and Rheumatoid Arthritis: What They Have in Common. Am. J. Med. 2020, 133, e91–e97. [Google Scholar] [CrossRef]
  118. Okumu, F.O.; Moore, S.J. Combining indoor residual spraying and insecticide-treated nets for malaria control in Africa: A review of possible outcomes and an outline of suggestions for the future. Malar. J. 2011, 10, 208. [Google Scholar] [CrossRef]
  119. Huang, Y.; Chen, Z.; Lan, J.; Zhang, L.; Chen, H.; Jiang, L.; Yu, H.; Liu, N.; Liao, C.; Han, Q. MDR49 coding for both P-glycoprotein and TMOF transporter functions in ivermectin resistance, trypsin activity inhibition, and fertility in the yellow fever mosquito, Aedes aegypti. Pestic. Biochem. Physiol. 2024, 201, 105899. [Google Scholar] [CrossRef]
  120. Lan, J.; Wang, Z.; Chen, Z.; Zhang, L.; Zhao, J.; Guan, Q.; Liao, C.; Liu, N.; Han, Q. Identification of the Aedes aegypti nAChR gene family and molecular target of spinosad. Pest Manag. Sci. 2021, 77, 1633–1641. [Google Scholar] [CrossRef]
  121. Severson, D.W.; Behura, S.K. Mosquito genomics: Progress and challenges. Annu. Rev. Entomol. 2012, 57, 143–166. [Google Scholar] [CrossRef] [PubMed]
  122. Viveiros-Rosa, S.G.; Regis, E.G.; Santos, W.C. Vector competence of Culex mosquitoes (Diptera: Culicidae) in Zika virus transmission: An integrative review. Rev. Panam. Salud. Publica. 2020, 44, e7. [Google Scholar] [CrossRef] [PubMed]
  123. Shaw, W.R.; Catteruccia, F. Vector biology meets disease control: Using basic research to fight vector-borne diseases. Nat. Microbiol. 2019, 4, 20–34. [Google Scholar] [CrossRef]
  124. Wang, H.; Shi, Y.; Wang, L.; Liu, S.; Wu, S.; Yang, Y.; Feyereisen, R.; Wu, Y. CYP6AE gene cluster knockout in Helicoverpa armigera reveals role in detoxification of phytochemicals and insecticides. Nat. Commun. 2018, 9, 4820. [Google Scholar] [CrossRef] [PubMed]
  125. Gc, G.; Parajuli, K.; Gautam, I.; Banjara, M.R.; Ghimire, P.; Rijal, K.R. Efficacy of Native Bacillus thuringiensis against Mosquito Vector. J. Nepal Health Res. Counc. 2024, 21, 479–485. [Google Scholar] [CrossRef]
  126. Dos Santos, C.A.M.; do Nascimento, J.; Gonçalves, K.C.; Smaniotto, G.; Zechin, L.D.; Ferreira, M.D.; Polanczyk, R.A. Compatibility of Bt biopesticides and adjuvants for Spodoptera frugiperda control. Sci. Rep. 2021, 11, 8. [Google Scholar] [CrossRef]
  127. Kang, R.J.; Li, R.; Mjengi, J.; Abbas, Z.; Song, Y.H.; Zhang, L. A tiny sample rapid visual detection technology for imidacloprid resistance in Aphis gossypii by CRISPR/Cas12a. Sci. Total. Environ. 2024, 951, 10. [Google Scholar] [CrossRef]
  128. Jin, M.; Shan, Y.; Peng, Y.; Wang, W.; Zhang, H.; Liu, K.; Heckel, D.G.; Wu, K.; Tabashnik, B.E.; Xiao, Y. Downregulation of a transcription factor associated with resistance to Bt toxin Vip3Aa in the invasive fall armyworm. Proc. Natl. Acad. Sci. USA 2023, 120, e2306932120. [Google Scholar] [CrossRef]
  129. Jin, W.; Zhai, Y.; Yang, Y.; Wu, Y.; Wang, X. Cadherin Protein Is Involved in the Action of Bacillus thuringiensis Cry1Ac Toxin in Ostrinia furnacalis. Toxins 2021, 13, 658. [Google Scholar] [CrossRef]
  130. Pacheco, I.D.; Walling, L.L.; Atkinson, P.W. Gene Editing and Genetic Control of Hemipteran Pests: Progress, Challenges and Perspectives. Front. Bioeng. Biotechnol. 2022, 10, 900785. [Google Scholar] [CrossRef]
  131. Mahmood, M.A.; Naqvi, R.Z.; Siddiqui, H.A.; Amin, I.; Mansoor, S. Current knowledge and implementations of Bemisia tabaci genomic technologies for sustainable control. J. Pest Sci. 2023, 96, 427–440. [Google Scholar] [CrossRef]
  132. Heu, C.C.; McCullough, F.M.; Luan, J.B.; Rasgon, J.L. CRISPR-Cas9-Based Genome Editing in the Silverleaf Whitefly (Bemisia tabaci). Cris. J. 2020, 3, 89–96. [Google Scholar] [CrossRef] [PubMed]
  133. Chen, Y.; Zhang, J.; Qin, X.W.; Liu, J.; Liu, Q.; Zhang, R.J. Sublethal Effects of VirtakoTM on Life Table Parameters and Wing Formation of the Brown Planthopper (Homoptera: Delphacidae). J. Entomol. Sci. 2014, 49, 21–29. [Google Scholar]
  134. Zhang, F.; Zhang, Y.C.; Yu, Z.T.; Zeng, B.; Sun, H.; Xie, Y.Q.; Zhu, K.Y.; Gao, C.F. The G932C mutation of chitin synthase 1 gene (CHS1) mediates buprofezin resistance as confirmed by CRISPR/Cas9-mediated knock-in approach in the brown planthopper, Nilaparvata lugens. Pestic. Biochem. Physiol. 2024, 202, 105953. [Google Scholar] [CrossRef]
  135. Nozaki, T.; Kobayashi, Y.; Shigenobu, S. First record of the cedar bark aphid, Cinara cedri cedri Mimeur, 1936 (Hemiptera: Aphidoidea) in Japan, and identification of infecting Wolbachia strains. BioInvasions Rec. 2022, 11, 900–911. [Google Scholar] [CrossRef]
  136. Atencia-Pineda, M.C.; García-Leal, J.; Diaz-Ortiz, D.; Pareja-Loaiza, P.; Pacheco-Lugo, L.; Hoyos-López, R.; Calderón-Rangel, A.; Fragozo-Castilla, P.; Gutiérrez-Rodríguez, S.M.; Flores, A.E.; et al. Susceptibility to organophosphate insecticides in Aedes aegypti (Diptera: Culicidae) from northern Colombia and associated resistance mechanisms. Parasites Vectors 2025, 18, 7. [Google Scholar] [CrossRef]
  137. Dos Santos Hage, R.; Bohm, B.C.; Casagrande, C.P.; Silva, S.C.M.; Soares, A.T.; Lima, J.V.; Bruhn, N.C.P.; Bruhn, F.R.P. Spatiotemporal expansion of dengue in Brazilian Amazon between 2001 and 2021. Sci. Rep. 2025, 15, 1032. [Google Scholar] [CrossRef] [PubMed]
  138. Apte, R.A.; Smidler, A.L.; Pai, J.J.; Chow, M.L.; Chen, S.L.; Mondal, A.; Sánchez, C.H.M.; Antoshechkin, I.; Marshall, J.M.; Akbari, O.S. Eliminating malaria vectors with precision-guided sterile males. Proc. Natl. Acad. Sci. USA 2024, 121, 12. [Google Scholar] [CrossRef]
  139. Kandul, N.P.; Liu, J.; Sanchez, C.H.M.; Wu, S.L.; Marsha, J.M.; Akbari, O.S. Transforming insect population control with precision guided sterile males with demonstration in flies. Nat. Commun. 2019, 10, 84. [Google Scholar] [CrossRef]
  140. Shirai, Y.; Piulachs, M.D.; Belles, X.; Daimon, T. DIPA-CRISPR is a simple and accessible method for insect gene editing. Cell Rep. Methods 2022, 2, 100215. [Google Scholar] [CrossRef]
  141. Zhang, X.; Singh, A.; Soriano Martinez, K.; Ferree, P.M. Direct Parental (DIPA) CRISPR in the jewel wasp, Nasonia vitripennis. G3 Genes Genomes Genet. 2024, 14, JKAE095. [Google Scholar] [CrossRef]
  142. Matsuda, N.; Takahashi, M.; Shirai, Y.; Hinomoto, N.; Daimon, T. Direct parental CRISPR gene editing in the predatory bug Orius strigicollis, a biocontrol agent against small arthropods. Pest Manag. Sci. 2024, 80, 5465–5472. [Google Scholar] [CrossRef] [PubMed]
  143. Kondo, K.; Tanaka, A.; Kunieda, T. Single-step generation of homozygous knockout/knock-in individuals in an extremotolerant parthenogenetic tardigrade using DIPA-CRISPR. PLoS Genet. 2024, 20, e1011298. [Google Scholar] [CrossRef] [PubMed]
  144. Zhang, M.; Gong, L.; Zhao, Y.; Ma, Y.; Long, G.; Guo, H.; Liu, X.; Hull, J.; Dewer, Y.; Yang, C.; et al. Efficient DIPA-CRISPR-mediated knockout of an eye pigment gene in the white-backed planthopper, Sogatella furcifera. Insect Sci. 2023, 31, 1015–1025. [Google Scholar] [CrossRef] [PubMed]
  145. Esvelt, K.M.; Smidler, A.L.; Catteruccia, F.; Church, G.M. Concerning RNA-guided gene drives for the alteration of wild populations. eLife 2014, 3, e03401. [Google Scholar] [CrossRef]
  146. Gouda, M.N.R.; Jeevan, H.; Shashank, H.G. CRISPR/Cas9: A cutting-edge solution for combatting the fall armyworm, podoptera frugiperda. Mol. Biol. Rep. 2024, 51, 13. [Google Scholar] [CrossRef]
  147. Lee, Y.; Oh, Y.; Lee, S.H. Recent advances in genome engineering by CRISPR technology. BMB Rep. 2024, 57, 12–18. [Google Scholar] [CrossRef]
  148. Zhao, F.; Ding, X.; Liu, Z.; Yan, X.; Chen, Y.; Jiang, Y.; Chen, S.; Wang, Y.; Kang, T.; Xie, C.; et al. Application of CRISPR/Cas9-based genome editing in ecotoxicology. Environ. Pollut. 2023, 336, 14. [Google Scholar] [CrossRef]
  149. Singh, S.; Rahangdale, S.; Pandita, S.; Saxena, G.; Upadhyay, S.K.; Mishra, G.; Verma, P.C. CRISPR/Cas9 for Insect Pests Management: A Comprehensive Review of Advances and Applications. Agriculture 2022, 12, 20. [Google Scholar] [CrossRef]
Insects 16 00345 g001
Figure 1. The architecture of ZFN, TALEN, and CRISPR/Cas systems. (A) The schematic of ZFN specifically recognizing and binding to DNA. Each DNA-recognition domain contains three zinc fingers, and each zinc finger structure is in direct contact with three bases. (B) The schematic of TALEN specifically recognizing and binding to DNA. The TALEN element is targeted and bound to specific DNA sites through a DNA-recognition module and then clipped at specific sites under the action of the Fok I nuclease. The RVD compositions are indicated in the dotted box (C) The schematic of cleavage by the Cas9 enzyme. The Cas9 enzyme recognizes the PAM (NGG) site and cleaves the target DNA sequence between the third and fourth bases near the PAM site. (D) The schematic of CRISPR site structure. The CRISPR/Cas system consists of CRISPR sequence elements and the Cas gene family. (E) The repair pathway of double-strand break (DSB) mediated by CRISPR system. The DSB induced by the Cas9/sgRNA complex can be repaired by non-homologous end joining (NHEJ) or homologous recombination (HR). This can result in small insertions or deletions at the target sites (left) and homologous repair with a desired template (right).
Figure 1. The architecture of ZFN, TALEN, and CRISPR/Cas systems. (A) The schematic of ZFN specifically recognizing and binding to DNA. Each DNA-recognition domain contains three zinc fingers, and each zinc finger structure is in direct contact with three bases. (B) The schematic of TALEN specifically recognizing and binding to DNA. The TALEN element is targeted and bound to specific DNA sites through a DNA-recognition module and then clipped at specific sites under the action of the Fok I nuclease. The RVD compositions are indicated in the dotted box (C) The schematic of cleavage by the Cas9 enzyme. The Cas9 enzyme recognizes the PAM (NGG) site and cleaves the target DNA sequence between the third and fourth bases near the PAM site. (D) The schematic of CRISPR site structure. The CRISPR/Cas system consists of CRISPR sequence elements and the Cas gene family. (E) The repair pathway of double-strand break (DSB) mediated by CRISPR system. The DSB induced by the Cas9/sgRNA complex can be repaired by non-homologous end joining (NHEJ) or homologous recombination (HR). This can result in small insertions or deletions at the target sites (left) and homologous repair with a desired template (right).
Insects 16 00345 g001
Insects 16 00345 g002
Figure 2. The application of CRISPR/Cas9 technology in insecticide resistance. Red circles represent single-gene knockout or knock-in, and blue circles represent multi-gene knockout. The orange arrow represents Diptera, the pink arrow represents Lepidoptera, and the light blue arrow represents Hemiptera. Diptera (Aa, Aedes aegypti; Ag, Anopheles gambiae; Cq, Culex quinquefasciatus; Dm, Drosophila melanogaster); Lepidoptera (Cs, Chilo suppressalis; Ha, Helicoverpa armigera; Hz, Helicoverpa zea; Of, Ostrinia furnacalis; Pg, Pectinophora gossypiella; Px, Plutella xylostella; Se, Spodoptera exigua; Sf, Spodoptera frugiperda; Ta, Tuta absoluta; Tn, Trichoplusia ni); and Hemiptera (Bt, Bemisia tabaci; Mp, Myzus persicae; Nl, Nilaparvata lugens). Sources Refs. [51,52,53,54,55,56,57,58,59,60,61,62,63,64,65,66,67,68,69,70,71,72,73,74,75,76,77,78,79,80,81,82,83,84,85,86,87,88,89,90,91,92,93,94,95,96,97,98,99,100,101,102,103,104,105,106,107,108,109,110,111,112,113].
Figure 2. The application of CRISPR/Cas9 technology in insecticide resistance. Red circles represent single-gene knockout or knock-in, and blue circles represent multi-gene knockout. The orange arrow represents Diptera, the pink arrow represents Lepidoptera, and the light blue arrow represents Hemiptera. Diptera (Aa, Aedes aegypti; Ag, Anopheles gambiae; Cq, Culex quinquefasciatus; Dm, Drosophila melanogaster); Lepidoptera (Cs, Chilo suppressalis; Ha, Helicoverpa armigera; Hz, Helicoverpa zea; Of, Ostrinia furnacalis; Pg, Pectinophora gossypiella; Px, Plutella xylostella; Se, Spodoptera exigua; Sf, Spodoptera frugiperda; Ta, Tuta absoluta; Tn, Trichoplusia ni); and Hemiptera (Bt, Bemisia tabaci; Mp, Myzus persicae; Nl, Nilaparvata lugens). Sources Refs. [51,52,53,54,55,56,57,58,59,60,61,62,63,64,65,66,67,68,69,70,71,72,73,74,75,76,77,78,79,80,81,82,83,84,85,86,87,88,89,90,91,92,93,94,95,96,97,98,99,100,101,102,103,104,105,106,107,108,109,110,111,112,113].
Insects 16 00345 g002
Table 1. Application of CRISPR/Cas9 system in insecticide resistance in insects.
Table 1. Application of CRISPR/Cas9 system in insecticide resistance in insects.
ClassificationSpeciesGenesMutation SummaryReferences
DipteraDrosophila melanogasterCyp6g1,
Cyp6g2		KnockoutmonogenicOnly high expression of Cyp6g1 contributes to imidacloprid resistance, while overexpression of Cyp6g2 can metabolize imidacloprid and produce resistance.Denecke + 2017 [51]
Cyp6g1KnockoutmonogenicThe Cyp6g1 gene is associated with the oxidative metabolism of insecticides but does not directly control the production of nitro-reducing metabolites.Fusetto + 2017 [52]
L1014Knock-inmonogenicThe CRISPR-based allele drive replaces the resistant kdr mutation with a susceptible wild-type counterpart.Kaduskar + 2022 [53]
Cyp6BQ23KnockoutmonogenicReduced pyrethroid affinity at the target site, delaying saturation while simultaneously extending the duration of P450-driven detoxificationSamantsidis + 2020 [54]
G275EKnock-inmonogenicThe G275E gene was associated with spinosad resistance using the CRISPR/Cas9 system.Zimmer + 2016 [55]
Dα6KnockoutmonogenicThe mechanism of nAChR’s response to three different chemical classes of insecticides was explored by CRISPR.Perry + 2021 [56]
I161V, G265A, F330YKnockoutmonogenicInsecticide susceptibility gene drives could be useful tools to control pest insects; however, problems with particularities of target loci and GDR will need to be overcome for them to be effective.Hernandes + 2024 [57]
VAChT Y49NKnock-inmonogenicThe GAL4/UAS system was combined with CRISPR technology to study the effect of VAChTY49N on CASPP resistance.Vernon + 2018 [58]
Mdr49, Mdr50,
Mdr65		KnockoutmonogenicDemonstrated the role of ABC transporter in insecticide tolerance of D. melanogasterDenecke + 2017 [59]
I276CKnockoutpolygenicThe intersubunit amino acids within the transmembrane M1 and M3 domains do form binding sites that are critical for the interaction of m-diamide and isoxazoline insecticides.Zhou + 2024 [60]
Aedes aegyptiAae-mALP, Aae-CadKnockoutmonogenicIn addition to Aae-Cad and Aae-mALP, other midgut membrane proteins are also involved in the Cry toxin receptor mode of Aspergillus aegypti.Pacheco + 2024 [61]
Culex quinquefasciatusActin5C, Rpl40, vasa, nanosKnockoutmonogenicThe gRNA scaffold variant improved the transgene efficiency of Culex mosquitoes.Feng + 2021 [62]
Anopheles gambiaeL1014FKnock-inmonogenicMosquitoes carrying the L1014F allele show an adaptive disadvantage in the homozygous state.Grigoraki + 2021 [63]
V402L, L995FKnock-inmonogenicIn some cases, the lower fitness costs associated with this kdr mutation may have a selective advantage over classical kdr.Williams + 2022 [64]
LepidopteraPlutella xylostellaABCC2, ABCC3KnockoutmonogenicTwo knockout strains were successfully constructed using the new CRISPR/Cas9 genome engineering system.Guo+ 2019 [65]
APN6, APN5, ABCC1KnockoutpolygenicReveals how P. xylostella adjusts MAPK phosphorylation in response to toxins and alters FTZ-F1 transcription factor binding to regulate the expression of Bt receptors or non-receptor paraloguesGuo + 2022 [66]
LepidopteraPlutella xylostellaAPN1, APN3aKnockoutmonogenicEvidence that the MAPK cascade response can be activated by enhanced upstream hormone signaling to counter Bt virulence in the diamondback moth.Guo + 2022 [67]
ABCC2, ABCC3
APN1, APN3a		KnockoutpolygenicBt toxins have multiple modes of action that can compensate for the loss of a single receptor.Sun D. + 2022 [68]
PxmALP, PxABCC2, PxABCC3KnockoutpolygenicReveals functional redundancy between ABC transporter proteins and PxmALPSun D. + 2024 [69]
PxABCC2,
PxABCC3		KnockoutpolygenicPxABCC2 and PxABCC3 are redundant or complementary.Liu + 2020 [70]
PxABCC2,
PxABCC3		KnockoutpolygenicThe value of using single-gene knockout and multi-gene knockout is emphasized.Zhao + 2020 [71]
PxABCB1KnockoutmonogenicPxABCB1 protects insects from avermectin insecticides; on the other hand, it promotes the toxic effects of Bt Cry1Ac toxin.Ouyang + 2024 [72]
PxMetAP1KnockoutmonogenicRevealed the important role of the MetAP gene in DBMBt toleranceYe + 2022 [73]
PxJHBPKnockoutmonogenicPxJHBP is a key gene in resistance to Cry1Ac and regulation of female reproduction.You + 2024 [74]
SE2KnockoutmonogenicAdoption of high-efficiency double sgRNA strategyGuo + 2023 [75]
miR-8545KnockoutmonogenicIncreased expression of microRNA (miR-8545) inhibits the newly discovered molting steroid degrading enzyme (PxGLD)Guo + 2024 [76]
I1056M/FKnockoutmonogenicWith the D. Melanogaster model, scientists were able to quickly observe phenotypic changes after gene editing, thus verifying the effect of the mutation.Douris et al. + 2016 [77]
PxJHEKnockoutmonogenicCRISPR/Cas9-induced m6A site-specific mutation PxJHE induces fitness costsGuo + 2024 [78]
nAChRα6KnockoutmonogenicEndogenous functional studies demonstrated the causal relationship of Pxα6 truncated mutations.Wang + 2020 [79]
V263IKnockoutmonogenicThe function of V263I mutation in PxGluCl was verified for the first timeSun + 2023 [80]
D472NKnockoutmonogenicThe homozygous D472N mutation in Rdl1 confers a low level of resistance to avermectin in P. xylostellaSun X. + 2023 [81]
I4790MKnock-inmonogenicThe functional role of PxRyR’s I4790M mutation in diamide resistance was confirmed for the first time.Wang X + 2020 [82]
LepidopteraPlutella xylostellaI4790KKnock-inmonogenicThe I4790K mutation reduces insecticide binding to receptorsJiang + 2024 [83]
Y4667D, I4758M, G4915EKnock-inmonogenicMultiple mutations in RyR confer resistance to diamides in clostridium-inhibiting bacteria.Huang + 2020 [84]
Helicoverpa armigeraHaCadKnockoutmonogenicHaCad provides strong reverse genetic evidence as a functional receptor of Cry1Ac.Wang + 2016 [85]
HaRdl-1, HaRdl-2KnockoutmonogenicHaRdl-1 and HaRdl-2 are important determinants of H. armigera sensitivity to three cyclodiene insecticides.Wang J. + 2020 [86]
Haα6, Haα7KnockoutmonogenicVariation in nAChRα6 was associated with high resistance of pests to spinosyns.Wang J. + 2020 [87]
HaABCA2KnockoutmonogenicThe midgut brush marginal membrane vesicles of knockout populations lost their ability to bind to Cry2Ab, but retained their ability to bind to Cry1Ac.Wang J. + 2017 [88]
HaABCC2, HaCADKnockoutmonogenicThe synergistic effect of CAD and ABCC2/ABCC3 significantly enhanced the resistance of H. armigera to Cry1Ac.Zhang D. + 2021 [89]
GSTKnockoutpolygenicThe complex changes in GST cluster expression enhanced resistance of field populations to the highly efficient pyrethroid.Jin + 2023 [90]
Pectinophora gossypiellaPgABCA2KnockoutmonogenicDemonstrated that destructive mutations lead to actual resistance to Cry2Ab and are associated with field resistanceFabrick + 2021 [91]
Helicoverpa zeaHzABCA2KnockoutmonogenicThe mutation of the HzABCA2 gene is a key factor leading to the development of resistance to Cry2Ab in Helicoverpa zea.Fabrick + 2022 [92]
Spodoptera frugiperdaCYP304F1KnockoutmonogenicCYP304F1 plays an important role in resistance to β-cypermethrin and chlorpyrifos.Xu + 2022 [93]
SfCHS2KnockoutpolygenicIdentified for the first time that the LTR retrotransposon Yaoer plays a pivotal role in the resistance mechanism against Vip3AaLiu + 2024 [94]
nAChRα6KnockoutpolygenicThe team used CRISPR to knockout Sfα6 in S. frugiperda, studying its role in spinosyn susceptibility.Shi + 2022 [95]
CYP9AKnockoutpolygenicCYP9A gene cluster knockout in S.exigua and S. frugiperdaShi + 2023 [96]
Trichoplusia niABCA1, ABCA2KnockoutmonogenicABCA2 is critical to the toxicity of Cry2Ab in T. ni.Yang X. + 2019 [97]
ABCC2Knock-inmonogenicInvestigated the association between ABCC2 and Trichoplusia ni resistanceMa X. + 2022 [98]
Spodoptera exiguaG4946EKnock-inmonogenicThe G4946E mutation in the RyR gene was shown to confer a high level of resistance to diamide insecticides.Zuo Y. + 2017 [99]
SeP-gpKnockoutmonogenicOverexpression of SeP-gp may lead to abamectin and EB resistance in S.exigua.Zuo Y. + 2018 [100]
Seα6KnockoutmonogenicProved the functional role of Seα6 in the treatment of spinosad and spinetoramZuo Y + 2020 [101]
G275EKnock-inmonogenicVerified the role of G275E mutation in S. exiguan AChRα6 in resistance to spinosynZuo Y. + 2022 [102]
Seα1KnockoutmonogenicSeα1 knockout results in the loss of functional transmembrane (TM)3 and TM4 elements.Wang Z. + 2023 [103]
Ostrinia furnacalisABCC2KnockoutmonogenicThe OfABCC2 protein may function as a receptor for Cry1Fa, enhancing its association with Cry1Fa toxin’s mode of action.Wang X. + 2020 [104]
ABCG4KnockoutmonogenicThe mutant exhibited enhanced Cry1 toxin resistance, impacting larval development and population.Gao + 2022 [105]
LepidopteraChilo suppressalisI4758M, Y4667D, Y4667C, Y4891FKnock-inmonogenicRevealed the role of RyR mutation in diamide resistance and how the mutation affects the binding affinity of different diamidesHuang + 2021 [106]
Y4667D, G4915E,
Y4667C, I4758M,
Y4891F		KnockoutpolygenicThe I4758M and Y4667C double mutations have higher tetraniliprole resistance than the single Y4667C mutation.Sun H. + 2024 [107]
Tuta absolutaF1845Y, V1848IKnockoutmonogenicThe V1848I and F1845Y mutations may have formed too large a side chain to affect metaflumizone binding.Samantsidis + 2019 [108]
G4946V, I4790MKnock-inmonogenicConfirmed the role of RyR mutations in diamide resistance and revealed how mutations affect the binding affinity of different diamidesDouris et al. + 2017 [109]
HemipteraNilaparvata lugensCYP6ER1KnockoutmonogenicCYP6ER1 activity is related to the structure of an insecticide.Zhang H. + 2023 [110]
NlCYP6CS1KnockoutmonogenicNl6CS1KO is similar to the wild type in development and longevity, but there are differences in survival, reproduction, and body weight.Zhang Y. + 2023 [111]
Bemisia tabaciA2083VKnockoutmonogenicB. tabaci is highly resistant to ketoenol insecticides, which is not related to metabolic resistance but is caused by A2083V mutation in the CT domain of ACC.Lueke + 2020 [112]
HemipteraMyzus persicaeR81TKnockoutmonogenicIntroduction of the R81T mutation in the black-bellied fly maggot using CRISPR/Cas9 results in enhanced resistance to neonicotinoid insecticides but reduced fitness.Homem + 2020 [113]
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Xu, Q.; Wang, M.; Zeng, J.; Sun, H.; Wei, X.; Jiang, H.; Shentu, X.; Sun, D. CRISPR/Cas Technology in Insect Insecticide Resistance. Insects 2025, 16, 345. https://doi.org/10.3390/insects16040345

AMA Style

Xu Q, Wang M, Zeng J, Sun H, Wei X, Jiang H, Shentu X, Sun D. CRISPR/Cas Technology in Insect Insecticide Resistance. Insects. 2025; 16(4):345. https://doi.org/10.3390/insects16040345

Chicago/Turabian Style

Xu, Qiuchen, Mingyun Wang, Jiahui Zeng, Hangzhen Sun, Xiaoqi Wei, Hui Jiang, Xuping Shentu, and Dan Sun. 2025. "CRISPR/Cas Technology in Insect Insecticide Resistance" Insects 16, no. 4: 345. https://doi.org/10.3390/insects16040345

APA Style

Xu, Q., Wang, M., Zeng, J., Sun, H., Wei, X., Jiang, H., Shentu, X., & Sun, D. (2025). CRISPR/Cas Technology in Insect Insecticide Resistance. Insects, 16(4), 345. https://doi.org/10.3390/insects16040345

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop