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Review

CRISPR-Cas Genome Editing for Insect Pest Stress Management in Crop Plants

by
Tasfia Tasnim Moon
1,
Ishrat Jahan Maliha
1,
Abdullah Al Moin Khan
2,
Moutoshi Chakraborty
2,
Md Sharaf Uddin
3,
Md Ruhul Amin
1 and
Tofazzal Islam
2,*
1
Department of Entomology, Bangabandhu Sheikh Mujibur Rahman Agricultural University (BSMRAU), Gazipur 1706, Bangladesh
2
Institute of Biotechnology and Genetic Engineering (IBGE), Bangabandhu Sheikh Mujibur Rahman Agricultural University, Gazipur 1706, Bangladesh
3
Department of Agroforestry and Environmental Science, Sylhet Agricultural University, Alurtol Road, Sylhet 3100, Bangladesh
*
Author to whom correspondence should be addressed.
Stresses 2022, 2(4), 493-514; https://doi.org/10.3390/stresses2040034
Submission received: 23 September 2022 / Revised: 30 November 2022 / Accepted: 1 December 2022 / Published: 7 December 2022
(This article belongs to the Special Issue Physiological and Molecular Mechanisms of Plant Stress Tolerance)
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Abstract

:
Global crop yield and food security are being threatened by phytophagous insects. Innovative methods are required to increase agricultural output while reducing reliance on hazardous synthetic insecticides. Using the revolutionary CRISPR-Cas technology to develop insect-resistant plants appears to be highly efficient at lowering production costs and increasing farm profitability. The genomes of both a model insect, Drosophila melanogaster, and major phytophagous insect genera, viz. Spodoptera, Helicoverpa, Nilaparvata, Locusta, Tribolium, Agrotis, etc., were successfully edited by the CRISPR-Cas toolkits. This new method, however, has the ability to alter an insect’s DNA in order to either induce a gene drive or overcome an insect’s tolerance to certain insecticides. The rapid progress in the methodologies of CRISPR technology and their diverse applications show a high promise in the development of insect-resistant plant varieties or other strategies for the sustainable management of insect pests to ensure food security. This paper reviewed and critically discussed the use of CRISPR-Cas genome-editing technology in long-term insect pest management. The emphasis of this review was on the prospective uses of the CRISPR-Cas system for insect stress management in crop production through the creation of genome-edited crop plants or insects. The potential and the difficulties of using CRISPR-Cas technology to reduce pest stress in crop plants were critically examined and discussed.

1. Introduction

Insects are the primary biotic stressors that pose a severe threat to crop loss globally due to their direct feeding behavior on crops and their ability to spread plant diseases [1]. It is estimated that one-fourth of crops is destroyed annually by insects [2]. New techniques for the management of phytophagous insect pests would contribute to the protection of crops and, thus, raise the yield of crops. The major insects that cause significant decreases in crop production are sap-sucking and chewing pests [3]. Recent advances in the molecular foundation of the interactions between insects, plants, and biotechnological techniques, such as genome editing, offer solutions to these problems [4]. Recently, it was discovered that designed nucleases have an enormous potential for genome editing in both plants and insects [5,6]. The application of genome-editing methods has dramatically grown over time. At the moment, CRISPR-Cas is the most widely used method of genome editing. [7,8]. Genome editing using the CRISPR-Cas system has proven to be successful in creating a variety of agronomic traits, including a long-lasting resistance to insect pests [5].
Genome editing, or gene editing, is a method of genetic manipulation that involves inserting, deleting, labeling, changing, or substituting DNA into the genome of a living being in order to generate a desired attribute [9]. The four main categories of sequence-specific nucleases in gene editing to date are mega nucleases, zinc finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated proteins (Cas) [10,11]. The newest and most sophisticated genome-editing technology is that of the CRISPR-Cas RNA-guided nucleases that were modified from bacterial innate immune mechanisms and derived from Type II CRISPR-Cas9 mechanisms [12]. In order to secure food supplies for the world’s expanding population and to fulfill Sustainable Development Goal 2 (zero hunger), contemporary agriculture practices using stress-resistant crops and genetically modified crops are given emphasis. The use of synthetic chemical insecticides to control insect pests during crop production is expensive and hazardous to humans and to the environment. Additionally, it has a bad impact on biodiversity and unintended insects. We have already observed the application of numerous insect resistance genes in genetically modified crops, such as Bacillus thuringiensis (Bt)-insecticidal crystal proteins (ICPs), which has had a significant influence on productivity and sustainability [13]. CRISPR-Cas gene editing is emerging as an effective tool for the development of insect-resistant plants to promote sustainable agriculture. The process of developing resistance against insects with this promising tool includes changing the effect or target interactions, knocking out host-susceptible genes, uncoupling the antagonistic action of defense hormones, and so on [14]. CRISPR-Cas gene editing has been effectively used over the past ten years to create insect-resistant plants and to modify a number of insects. This approach has demonstrated great promise for increasing crop output through the sustainable management of insect pests. Insect pest control in agriculture could benefit from the development of genome-edited agricultural plants and insects. This review aimed to summarize the recent developments in the use of CRISPR-Cas in the generation of insect-resistant plants as well as in the application of this revolutionary technology to the modification of the genomes of phytophagous insects. We also discussed the difficulties and the potential use of the CRISPR-Cas toolbox for the sustainable management of insect pests.

2. CRISPR-Cas System and Its Mechanism of Genome Editing

CRISPR is the acronym for clustered regularly interspaced short palindromic repeats, and Cas is the CRISPR-associated protein. It is a natural defense mechanism found in the genomes of prokaryotic organisms such as bacteria (45%) and archaea (84%) [15]. It is a gene-editing technology that targets a specific DNA section, makes a precise cut at the target site, and makes the gene nonfunctional or replaces one version of the gene with another. It was invented by Jennifer Doudna, Emmanuelle Charpentier, and Fang Zhang in 2012, and the former two scientists were awarded the Nobel Prize in Chemistry in 2020 [16]. This is the most current technology for gene editing. This CRISPR-Cas9 editing technology is categorized into type I, II, and III, and, in the type II systems, the Cas9 nuclease requires DNA that matches a single RNA guide (sgRNA) [17]. The two primary components of the CRISPR-Cas system are the guide RNA and the Cas protein. The Cas9 protein, a nuclease enzyme that is commonly referred to as molecular scissors, is responsible for cutting the DNA. Guide-RNAs are molecules that direct Cas9 to the chosen spot in the genome, where it will remove the existing sequence and replace it with the new one [18]. It has recently become a very effective, fast, and speedy genome-editing tool [19,20]. This CRISPR-Cas technology can now be used to edit multiple genes at a time or even a single base, also known as epigenetic editing.
Several CRISPR-Cas applications have been shown to change the DNA sequences of insect or plant genomes [21,22,23]. Streptococcus pyogenes (Sp) is the source of the Cas9 protein that is currently most frequently employed [24]. In this procedure, a Cas9-protein-associated single-guide RNA (sgRNA) cleaves a particular target DNA region next to a protospacer adjacent motif (PAM), triggering the cellular DNA’s repair system to create a double-strand break (DSB). Without the homologous repair template, error-prone non-homologous end-joining (NHEJ) pathways are activated, resulting in spontaneous insertions/deletions or even replacements at the DSB site, which typically disrupt gene function. On the other hand, error-free homology-directed repair (HDR) mechanisms are activated, leading to mutations that undertake precise gene alterations, including knock-ins, knockouts, or mutations, if donor DNA templates are available that are similar to the sequence surrounding the DSB site [25]. The NHEJ and HDR mechanisms have currently been successfully co-opted for genome editing in a variety of insects and plants [14,22,26]. After successful genome modification, the CRISPR-Cas construct is delivered into plant cells through Agrobacterium-mediated or particle-bombardment-mediated transformation methods and into the embryos of insects through microinjection, transfusion, or electroporation-mediated transformation methods for the regeneration of transgenic species with desired characteristics [14,22,27]. The workflow of CRISPR-Cas genome editing in plants and insects is briefly illustrated in Figure 1.

3. CRISPR-Cas Genome Editing in Agriculture for the Management of Insect Pests

Biotechnology performs a crucial role in the control of insect pests to protect crops and improves yields in areas ranging from breeding for pest resistance to the genetically modified introgression of new genes [28]. The use of genome-editing techniques to create insect-resistant plants is still in its early stages. By manipulating the genes of both plants and insects, genome editing can be used to manage insect populations. The insect pests of crops can be controlled by inducing sterility in insect pests, interrupting pesticide resistance, or creating de novo resistance if adequate R-genes are lacking. Using CRISPR-Cas9 genome-editing technology, novel research is being done to modify insects to prevent them from feeding on and injuring plants and to modify plants to increase their efficacy in repelling insects [22,29]. In this respect, the genome-editing platform has offered a new opportunity for generating designer plants, especially in circumstances where a targeted deletion is likely to produce elite and superior characteristics or to trigger a gene drive to selectively spread mutations contributing to the lethality of female insect populations.
The agricultural biotechnology sector has been threatened with the problem of insect resistance to the Bt trait; thus, biotech companies are searching for a novel, economically viable, and environmentally responsible solution to the problem. In the biotech sector, CRISPR-Cas9 gene editing has emerged as the leading method for the control of insect pests [30]. In order to successfully alter a gene’s function, genome-editing technology actually leverages the cell’s own internal processes. Genome editing makes sure that the DNA sequence of a specific target genome is altered via the addition, deletion, and/or substitution of DNA bases [31].

3.1. CRISPR-Cas Genome Editing in Insects

In agriculture, CRISPR-Cas can be employed for crop protection through insect pest management. The genome editing of insects can be carried out successfully witha two-step technique involving the alteration of target DNA in insects and their eventual release into nature [32]. One of the earliest documented uses of the CRISPR-Cas system in insects was in Drosophila fruit flies, where effective modifications of the yellow gene were made [33].
The BmBLOS2 gene was the focus of another reported successful application of this method in silkworms [34], which was followed by several successful applications. In a case study by Garczynski et al. [35], the codling moth genome was edited using CRISPR-Cas gene-editing technology in order to alter the viability and production of eggs by targeting a particular gene (CpomOR1). Worldwide, the codling moth is a significant pest to pome fruit. As a member of the pheromone receptor subfamily, the CpomOR1 gene product is an odorant receptor. In the early-stage eggs of codling moths, single-guide RNAs (sgRNAs) were created to target the nucleotides of the CpomOR1 gene. It was discovered that alterations, including insertions and deletions, were successfully introduced. By mating males with females who had CpomOR1 gene alterations, the study tried to produce stable populations of edited codling moths by raising the young moths to adulthood. It was discovered that the modified females’ fecundity and fertility were compromised, causing them to produce non-viable eggs. The result was the regulation of fruit pomes by the insects. However, it is still unclear exactly how CpomOR1 affects the fertility and reproduction of codling moths. In another case, it was claimed that the migratory locust underwent a targeted heritable mutation as a result of the CRISPR-Cas technique. Locusts are dangerous agricultural pests that have an impact on a wide variety of crop plants. Their swarming behavior can result in very serious crop damage over large areas all at once, frequently leading to significant financial loss. The Li et al. [36] study involved the engineering of the guide-target RNA’s sequence to prevent the odor receptor co-receptor gene from being expressed (Orco). Orco gene mutants were shown to have defective electrophysiological reactions to several odors, resulting in mutant locusts lacking their attraction to aggregation pheromones under crowding circumstances.
Although the transgenic Bt technology is well established and widely utilized, the development of insect resistance to Bt insecticidal proteins (ICPs) has become a significant concern. In order to avoid this, efforts are being made to build receptors in a way that will enable effective resistance management. By altering the Helicoverpa armigera genome, it is possible to successfully knock down the Cadherin receptors that are functionally connected to Cry1Ac toxin tolerance [37]. A base replacement in the encoding genes of the mid-intestinal receptor demonstrated how the genome of insects can change their resistance to insect pests. Modifying Cry protein binding receptors can be used to edit insect genomes to decrease plant vulnerability. Unique detoxifying enzymes produced by insects are crucial for resolving the chemical defense responses in many plant species. A possible alternative would be to focus on polyphagous bugs’ detoxifying genes. Insect susceptibility resulted from targeting and deleting insecticidal detoxifying genes, such as gossypol-inducing cytochrome P450 [38]. The polyphagous insect H. armigera’s susceptibility to phytotoxins was revealed with the CRISPR-Cas-mediated deletion of the CYP6AE gene cluster, which also made crops resistant to insects and showed the importance of these enzymes in the detoxification of several toxic phytochemicals [39]. The most long-lasting answer has consistently been this one.
The modification of target genes that can prevent chemical contact and mating pair recognition, which are crucial for efficient interactions between plants and insects, is another method to control insects using CRISPR-Cas. The olfactory receptors (ORs) in insects are crucial for the identification of host plants and mating pair odorants. The Or83b gene mutation in Drosophila prevented the host from being detected [40]. Similar to this, the CRISPR-Cas method’s deletion of the Orco gene from Spodopthera litura affected its choice of a mating partner and host plant [32]. Implementing such technology would be a smart move to keep insects away from plants and to prevent pest damage. In insects, female adults release pheromones that males pick up on. Males select mature females based on their pheromone cues. A CRISPR-Cas9-based odorant receptor 16 (OR16) knockout in H. armigera prevented males from detecting pheromone signals and prevented mating with immature females, which led to the dumping of infertile eggs and helped in controlling insects [41]. Another strategy for the control of insects is to use CRISPR-Cas9 to remove growth genes, such as the abd-A (Abdominal A) gene, from a variety of insects, including Spodoptera litura [42], Spodoptera frugiperda [21], and Plutellaxy lostella [43], which resulted inabnormal gonads, disarmed prolegs, and the lack of body segment functions. The CRISPR-Cas9 technology was used to modify numerous other genes in a variety of insect pests. In Drosophila melanogaster, the LUBEL, Scsa, and Kdr genes were knocked out through CRISPR-Cas to limit normal growth and insecticide resistance [44]. Additionally, Chitin synthase 1 and nicotinic acetylcholine receptor α6 were replaced in order to limit insect population growth and insecticide resistance [45,46]. Scsa and Kdr genes were also knocked out for insecticide resistance [47,48]. In the case of Spodopteraexigua, the ryanodine receptor was substituted to control the insect population and its resistance to various insecticides [49], and the CYP9A186 gene, a-6-nicotinic acetylcholine receptor (nAchR), and P-glycoprotein gene were knocked out to make the species susceptible to emamectin benzoate (EB) [50], and to increase its susceptibility to abamectin and emamectin benzoate [51]. Genome editing of the SfABCC2 gene of S. frugiperda conferred resistance to the Cry1F toxin of B. thuringiensis [52] and two ABC transporters were differentially implicated in the toxicity of the two Bacillus thuringiensis Cry1 toxins of the invasive crop insect S. frugiperda [53]. Additionally, in S. frugiperda, the deletion of the ABCB1 gene increased its susceptibility to emamectin benzoate, beta-cypermethrin, and chlorantraniliprole [54].To create resistance in Helicoverpa armigera to cry2Aa and cry2Ab, the HaABCA2 gene was knocked out with CRISPR-Cas [55]. The nAChR6 gene was knocked out in Plutellaxy lostella to render it resistant to spinosad [56]. Dendrolimus punctatus had the DpWnt-1 gene knocked out, which caused defects in appendage development and anterior segmentation [57]. Cinnabar and White genes were altered to change the eye pigmentation in Bemisia tabaci and Nilaparvata lugens [58,59]. Malformations in embryonic development were caused by the CRISPR-Cas-9 disruption of the White and paired genes in Ceratitis capitata [60] (Table 1).
Table
Table 1. CRISPR-Cas genome editing in insects for insect pest management.
Table 1. CRISPR-Cas genome editing in insects for insect pest management.
Name of InsectTarget GeneEditingOutcome
Drosophila
melanogaster		YellowKnockout,
Knock-In		Generated designer flies [33]
Rosy and DSH3PX1Knockout,
Knock-In		Executed efficient and complex genomic manipulations [33]
LUBELKnockoutReduced survival rate [44]
Chitin synthase 1SubstitutionControlled insect population and resistance to various insecticides [45]
Nicotinic acetylcholine receptor α6SubstitutionControlled insect population and resistance to various insecticides [46]
ScsaKnockoutReduced normal growth [47]
KdrKnockoutReduce insecticide resistance [48]
Ast, Eh, capa, Ccap, Crz, npf,
Mip, mir-219, mir-315, and white		KnockoutTargeted mutagenesis [61]
YellowKnockoutEffectively targeted mutagenesis [62]
Yellow and whiteKnockoutHighly efficient and varied genome-editing efficiencies [63]
Yellow and rosyKnockout,
Knock-In		First report using the
CRISPR/Cas9 system to mediate
efficient genome engineering in
Drosophila [64]
AlkKnockoutEstablishing mutations [65]
TpnCKnockoutConfirmed that the myofibril assembly
wasrelated to TpnC gene [66]
WntlessKnockoutAmplified the cleavage
Efficiency [67]
Yellow, white, and tanKnock-InAttaining single or multiple allelic substitutions [68]
Act5C, lig4, and mus308Knockout,
Knock-In		Genome editing in Drosophila S2 cells [69]
Mod(mdg4)KnockoutValidation of a functional gene involved
in trans-splicing that influenced the
development in flies [70]
FdlKnockoutAnalyzing or manipulating protein
glycosylation pathways [71]
Chameau, CG4221, and CG5961 mRNAKnock-InA problem associated with “ends-in”
recombination was resolved [72]
ClampKnockoutThe expression of a sex-specific gene
was regulated [73]
Dα6Knock-InResistance to spinosad [46]
YellowKnock-InHeterozygous recessive mutation was
converted to homozygous loss of
function [74]
Ebony, yellow, wg, wls, Lis1, and SeKnockout,
Knock-In		Non-transgenic individuals exhibited less efficient knock-in than transgenic individuals did [75]
Ebony, yellow, and whiteKnockout,
Knock-In		Enhanced efficiency of gene targeting [76]
Ebony, yellow, and vermilionKnockout,
Knock-In 		Donor template and sgRNA plasmids
were injected into Cas9 transgenic
embryos in Drosophila [77]
White and piwiKnockout,
Knock-In		Prevented off-target effects during the
generation of indel mutants [77]
SalmKnock-InFlexible modification of fly genome [78]
Yellow,
notch, bam, nos, ms(3)k81, and cid		KnockoutTemporally and spatially inhibited
gene expression [79]
Ms(3)k81, white, and yellowKnockout,
Knock-In		CRISPR-mediated genome editing was
shown in Drosophila [80]
EGFP and mRFPKnockoutInduction of mutations [81]
Ebony, yellow, wingless, and wntKnockout,
Knock-In		Different patterns of expression [82]
Drosophila
subobscura		Yellow and whiteKnockoutGene functions were analyzed in a
non-model Drosophila species [83]
Drosophila suzukiiWhite (w) and sex lethal (Sxl)KnockoutControlled insect population and resistance to various insecticides [27]
DsRed
(red fluorescence protein)		Knock-InStudied sexing and monitoring [84]
White (w-)KnockoutAbsence of mating and copulation
failure [85]
Spodoptera exiguaSeα6KnockoutResistance to insecticides [23]
Ryanodine receptorSubstitutionControlled insect population and resistance to various insecticides [49]
CYP9A186 geneKnockoutSusceptibility to emamectin benzoate (EB) [50]
P-glycoprotein geneKnockoutSusceptibility to abamectin and emamectin benzoate [51]
a-6-nicotinic acetylcholine
receptor (nAchR)		KnockoutResistance to spinosyn insecticides [23,65,66]
Spodoptera littoralisOrcoKnockoutReduced survival rate [32]
Spodoptera
litura		Abdominal-A (slabd-A)KnockoutDefected body segmentation and pigmentation [42]
SlitPBP3KnockoutDestroyed pest insect mating [86]
SlitBLOS2KnockoutColoration of the integuments, a marker gene for functional studies and
pest control strategies [87]
Spodoptera frugiperdaSfabd-AIndelDefected body segmentation [21]
BLOS2E93
TO		KnockoutDeveloped mutants [88]
SfABCC2EditResistance to Cry1F toxin of Bacillus thuringiensis [52]
ABC transporters Toxicity of two Bacillus thuringiensis Cry1 toxins to the pest [53]
ABCB1KnockoutSusceptibility to emamectinbenzoate, beta-cypermethrin and chlorantraniliprole [54]
Helicoverpa armigeranAchRKnockoutResistance to insecticides [23]
a-6-nicotinic acetylcholine
receptor (nAchR)		KnockoutResistance to spinosyn [23,68,69]
HaCadKnockoutResistance to Bttoxin Cry1Ac [37]
Cluster of nine P450 genesKnockoutIdentification of the key players in
the insecticide metabolism [39]
CYP6AEKnockoutRegulation of detoxification enzymes [39]
OR16KnockoutDestroyed pest insect mating [41]
TetraspaninKnockoutResistance to Bt toxin cry1Ac [89]
HaABCA2KnockoutResistance to cry2Aa and cry2Ab [55]
White, ok, brown, and scarletKnockoutDifferential distribution of eye pigments that are helpful in
elucidation of biosynthetic pathway [90]
NPC1bKnockoutNPC1b wasvital for growth and
dietary cholesterol uptake [91]
HelicoverpapunctigeraHaABCA2DeletionResistance to both cry2Aa and cry2Ab [55]
Plutellaxy lostellaAbdominal-AKnockoutDefected body segmentation [43]
Pxabd-AKnockoutProviding novel ideas for pest management [43].
PxCHS1KnockoutDescribed the resistance management strategies of major agricultural pests [45]
PxABCC2
PxABCC3		KnockoutResistance to cry1 Ac protoxin [92]
nAChRα6KnockoutResistance to spinosad [56]
LWKnockoutWeaker phototaxis and reduced locomotion [93]
PxdsxKnockoutAltered expression of sex-biased genes [94]
Dendrolimus
punctatus		DpWnt-1KnockoutDefected anterior segmentation and appendage development [57]
Bemisia
tabaci		WhiteEditAltered eye pigmentation [58]
Nilaparvata
lugens		Cinnabar and whiteEditAltered eye pigmentation [59]
Nl-cn and
Nl-w		KnockoutPaved a path for gene-function interrogation [59]
Ceratitis
capitata		White eye (we)
and paired gene (Ccprd)		KnockoutEmbryonic developmental malformations [60]
eGFP_gRNA2,
eGFP_gRNA2, 1 mM Scr7, and
eGFP_gRNA2b–Cas9 complexes
with ssODN_BFP
donor templates		Knock-InConversion of eGFP to BFP [95]
Bactrocera dorsalisWhite and transformerKnockoutVarious phenotypic effects [96]
Anastrepha ludensAstra-2KnockoutThe mutation caused sterility [97]
Locust amigratoriaOrcoKnockoutGenerated lossoffunction for the management of insect pests [36]
OfAgo1KnockoutCuticle disruption [98]
Cydia pomonellaCpomOR1Knockout,
Knock-In		Affected egg production and viability [35]
Tetranychus
urticae		Phytoene desaturaseKnockoutFunctional studies [99]
PSSTKnockoutPyridaben resistance [100]
Leptinotarsa decemlineataVestigial gene (vest)KnockoutFunctionally characterized vest gene and mutagenesis [101]
Euschistus herosAbnormal wing disc (awd),
tyrosine hydroxylase (th), and
yellow (yel)		Knockdown and
knockout		Managing insect pests [102]
Diaphorina
Citri, Homalodisca
vitripennis, and
Bemisia argentifolii		Thioredoxin and vermillionKnockoutProtected food crops from different pathogens and insect vectors [103]
Diaphorina citriACP-TRX-2KnockoutInnovative breakthrough in gene editing [104]
Mythimna
separata		NPC1bKnockoutHampered nutrient absorption [105]
HyphantriacuneaHcdsxKnockoutSex-specific sterility [106]
Agrotis ipsilonAiTHKnockoutNarrowing in the eggshell [107]
Danaus plexippusClkKnockoutDefined the role of the clk gene in the control of migration behavior [108]
Bombyx moriBmBLOS2KnockoutGenerated designer flies [34]
BmOrcoKnockoutImpaired olfactory sensitivity [109]
Tribolium castaneumEGFPKnockout,
Knock-In		Controlled insect pests and created resistance to insecticides [110]
Tribolium E-cadherinKnockoutKnockout study [110]
Gryllus
bimaculatus		Dop1KnockoutDestroyed appetitive reinforcement [111]
Rhopalosiphum
padi		ß-1-3glucanaseKnockoutReduced callose deposition in maize [112]
Ostrinia
furnacalis		ABCC2KnockoutResistance to Btcry1Fa toxin [113,114]

3.2. CRISPR-Cas-Mediated Gene Drive in Insect Pest Management

Genome editing using CRISPR-Cas creates a gene drive that is effective enough to propagate the changed genes across generations until they are released for mating. A gene drive is a technique for the rapid distribution of altered genes throughout an insect species’ entire population. Gene drives based on CRISPR-Cas may cause sterility or mortality in targeted insect species due to gene disruption, which ultimately leads to a population collapse and even elimination due to severe recessive lethal changes [115]. A species will completely disappear as a result of this over the course of 15–20 generations. By selectively harming the X chromosome, the gene drive will alter the male sex ratio. This causes the Y chromosome to be more common in the most viable sperms, resulting in a greater proportion of male progeny and a progressive decline in the number of females [115]. Therefore, releasing insect strains with undesirable features, including lethality, infertility, a biased sex ratio, insecticidal sensitivity, etc., is a successful method for insect pest control. For instance, it should be assumed that the Bt resistance management in H. armigera is a sustainable method since, in this case, gene deletion would only affect the species of H. armigera that is resistant to Bt toxins [89].

3.3. CRISPR-Cas Technology in Genome Editing of Crop Plants

Technologies such as CRISPR-Cas can improve plant quality to preserve crops and help them survive specific biotic and abiotic challenges [6,113]. Maintaining healthy plants is a part of the Integrated Pest Management Program because insects are drawn to unhealthy, diseased plants. Plants can be modified using CRISPR-Cas systems so that they produce or do not produce particular enzymes that can deter insect pests from coming into contact with the plant or can attract specific insect predators to feed on the bug species that are attacking the plant [116]. The process of genome editing is quickly increasing its potential and its chances for giving insect resistance traits to crop plants. The lack of a clearly defined source of resistance in the gene pool, however, has led to less research on altering plants for pest management. The goal of several efforts in order to alleviate this bottlenecking is to collect genes from uncharacterized crop plant accessions and wild relatives. However, due to poorly understood resistance characteristic genetics in uncharacterized accessions, significant advances could not be made [117]. On the other hand, a transgenic method was used to introduce insect resistance genes into crops from more remote origins, such as the Bt genes from bacterial sources. These transgenic plant species, however, encountered severe political, moral, and social opposition because of a lack of scientific understanding [118]. In this situation, the main challenge in modern agriculture is to develop an environmentally sound breeding strategy for crops that can accomplish two breeding objectives: the production of de novo tolerance in the absence of the proper R-genes and the tracking of the dynamics of pests by destroying insecticide resistance, killing, or inducing insect sterility. Any insect will choose to lay eggs on the host plant if feed is available for its young. Plant volatile blends are combinations of volatiles that serve as cues for insects to select hosts and oviposition sites. Insects use their highly adaptable olfactory systems to detect suitable plants to serve as hosts by detecting volatile secondary chemicals in plants. According to the research done by Beale et al., altering volatile mixtures through genome editing can kill insects on host plants while making the plants resistant to them. When plants become infested with aphids, the sesquiterpene hydrocarbon (E)-β-farnesene (Eβf) is released, which reduces the populations of other hosts’ ability to eat while luring Diaeretiella rapae, a parasitic wasp that has been shown to dominate the aphid population in transgenic plants [119]. The genetic engineering of plant volatile blends may be a different strategy for insect management. However, care should be made to ensure that the change does not have a negative impact on the species of beneficial insects.
It is also possible to enhance the host’s immunity to pests by editing important plant immunity genes, such as genes regulating the target’s interactions with insect effectors and resistance genes (R-genes). Although S-genes make plants vulnerable to stress, R genes evaluate a plant’s susceptibility to insect pests and diseases [120]. The editing of R and S genes for the development of insect resistance in plant species is emerging as a dependable method. Due totheir growth, immunity, and behaviors that have been observed in rice, insects are known to be dependent on important chemical components contained in plants [22]. Genetic engineering in plants has been demonstrated in insect pest resistance by knocking off the S-genes ofthe plants. Tryptamine 5-hydroxylase encoding CYP71A1 gene deletion using CRISPR-Cas caused tryptamine’s conversion to serotonin in plants, which reduced plant hopper growth. Rice was alteredby Lu et al. [22] using the CRISPR-Cas9 technology to make it resistant to the striped stem borer and the brown plant hopper (Nilaparvat alugens) (Chilo suppressalis). The simultaneous deletion of two endogenous phytoene dehydrogenase (PDS) genes in P. tomentosa Carr., PtoPDS1, and PtoPDS2 using the CRISPR-Cas9 technique resulted in the effective generation of endogenous gene mutations in the Populus [121,122]. By enhancing their endogenous defenses, CRISPR-Cas genome-editing techniques also made it possible to increase the population’s resistance to insects. The golden promise barley variety’s two beta-1-3 glucanase genes were altered withCRISPR-Cas9, which reduced the amount of callus that formed in sieve tubes. Therefore, the aphid Rhopalosip humpadi could not access the phloem sap, which adversely affected its growth as well as disrupted its predilection for particular hosts [63]. On the basis of a plant’s outward appearance, insects can also recognize and target certain plants. It has been found that variations in plant color can influence an insect’s host preferences. This was confirmed in red-leaf tobacco made by altering the anthocyanin pathway. By changing the color of the leaf, gene editing for insect pest tolerance in plants was demonstrated. This prevented the insect from recognizing the host plant. The red color of the leaves was the result of an excess of anthocyanin coloring. The Helicoverpa armigera and Spodoptera litura were discouraged by this color change [123]. This study demonstrated that CRISPR-based editing for pest management, where the insects are unable to recognize the host plant, may be resolved by altering the anthocyanin pathway. According to Li et al., the GmCDPK38 mutant with the Hap3 deletion in soybeans showed significant resistance to common cutworms [124]. Additionally, the GmUGT gene deletions of 1bp and 33bp were made in soybeans to improve their resistance to S. litura and H. armigera [125] (Table 2).
Table
Table 2. CRISPR-Cas genome editing in crops for insect pest management.
Table 2. CRISPR-Cas genome editing in crops for insect pest management.
Name of CropsTarget GeneEditingOutcome
BarleyBeta-1-3 glucanaseAlterationResistance to aphid infestation [112]
Paulownia tomentosaPtoPDS1 and PtoPDS2DeletionEnhanced endogenous defenses and increased resistance to insects [121,122].
TobaccoAnthocyanin pathwayAlterationDiscouraged insect attack [123]
SoybeanGmCDPK38KnockoutResistance to common cutworm [124]
GmUGT1bp and 33bp deletionEnhanced resistance to Helicoverpaarmigera and Spodopteralitura [125]
Cry 8 like Resistance to Coleopteran-Holtrichiapanallele [126]
Solanum pimpinellifoliumSix different genesEditingResistance to insect pests [127].
RiceOsCYP71A1DeletionResistance to the striped stem borer and the brown plant hopper [22].
Cry2ATransgeneResistance to leaf folder [128]
Mannose-specific GNATransgeneResistance toBPH (Nilaparvata lugens) and hemipteran pest [127]
Cry1AC + ASALTransgeneResistance to yellow stem borer, leaf folder, and BPH [129]
Cry1Ab + vip3ATransgeneResistance toAsian stem borer and rice leaf folder [130]
Cry2AX1TransgeneResistance to rice leaf folder [131]
Cry2Aa + cry1CaTransgeneResistance to Chilo suppressalis
[132]
Cry2AX1
(cry2Aa + cry2Ac)		TransgeneResistance to Lepidopteran pest [133]
CottonCry2AX1TransgeneResistance to H. armigera [134]
Cry2Ab, cry1F, and cry1ACTransgeneResistance to Lepidopteran pest
H. armigera, and S. litura [135]
Cry1AC and cry2AbTransgeneResistance to S. litura [136]
Cry2AXTransgeneResistance to H. armigera [137]
Cry1AaTransgeneResistance to Anthamous grandis [138]
Cry1AbTransgeneResistance to Heliothis [139]
Cry1Ab+ NptIITransgeneResistance to H. armigera [140]
Vip3AcAaa
(vip3Aa1 + vip3Ac1)		TransgeneResistance to Lepidopteran [141]
Vip3A + cry1AbTransgeneResistance to Heliothis. zea and H. virescens [142]
Insect gut-binding lectin from
Sclerotium rolfsii		TransgeneResistance to chewing and sucking pest [3]
Cry51Aa2TransgeneResistance to Lygus species
[143]
Cry1Be + cry1FaTransgeneResistance to S. litura and O. nubialis
[144]
Maize Cry1Ab/cry2AjTransgeneResistance to S. exigua and Harmonia axyridis [145]
Mustard (Brassica juncea)Lectin protease protein(lentil lectin-LL CPPI)TransgeneResistance toaphid [146]
Colocasia esculenta tuber agglutinin
(CEA) + GNA		TransgeneResistance tomustard aphid (Lipaphis erysimi) [147]
SugarcaneCry2Aa+ cry1Ca
Cry1Ab + cry1Ac		TransgeneResistance to shoot borer [148]
Vip3ATransgeneResistance to sugarcane stem borer (Chilo infuscatellus) [149]
Potato Hv1a/GNATransgeneResistance to peach potato aphids and grain aphids [150]
Galanthus nivalis agglutinin (GNA)TransgeneResistance to aphids [151]
Wheat Pinelliapedatisecta agglutinin (PPA)Transgene Resistance to aphids [152]
CowpeaArcl on APA locus of
Phaeselous vulgaris		Transgene Resistance to bruchids [153]
Vip3Ba1TransgeneResistance to legume pod borer (Maruca vitrata) [154]
Pigeon peaCry2AaTransgene Resistance to H. armigera [155]
Cry2AaTransgeneResistan topod borer—H. armigera [156]
Cry1AC, cry2AaTransgeneResistance to H. armigera [157]
ChickpeaCryIIAaTransgeneResistance to pod borer [158]
TomatoCry1AcTransgeneResistance to Tuta Absoluta—tomato leaf miner [159]
Remusatia vivipara (rvl 1) and Sclerotiumrolfsii(srl 1)TransgeneResistance to root knot nematode (Meloidogyne incognita) [160]
Cry1AbTransgeneResistance to T. absoluta [161]
CastorCry1ACTransgeneResistance to Lepidoteran—Achaea Janata and S. litura [162]
Sweet potatoCry1AaTransgeneResistance to S. litura [163]

3.4. Utilization of Crop Wild Relatives in Insect Resistance by CRISPR-Cas Technology

The insertion of foreign genes into plants is one of the key regulatory problems associated with transgenics that can be overcome with gene editing. The cultivated crops’ forebears and close relatives, known as crop wild relatives (CWRs), are robust to biotic and abiotic stress but have low yields. After domesticating wild species and breeding plants, however, the cultivable germplasms and crops had large yields and could meet other human needs, but they could not withstand insect assault. Using CRISPR-Cas9 genome editing, we can effectively delete or modify the genes that cause insect susceptibility, or we can introduce unique features from CWRs to the cultivated species to create new cultivars that are insect-resistant [120].
Two steps can be taken to implement this. First, the de novo domestication of crops with insect-resistant wild cousins can be implemented. Gene-editing techniques can be used to alter the desired agronomic traits that are caused by genes. There is evidence that the wild tomato Solanum pimpinellifolium is resistant to spider mites and other arthropod insect pests [164]. The multiplex CRISPR-Cas editing of six different genes in S. pimpinellifolium resulted in the production of a high-yielding tomato with insect and pest tolerance in a single generation [165]. This method, based on a plant’s properties and molecular pathways, can be carefully applied to other CWRs. The de novo domestication of CWRs may, therefore, be a ground-breaking method for the development of crops with improved characteristics.
Second, using genes found in CWRs that are insect-resistant, the genome can alter the cultivated crops. By altering the genomes of cultivated crops to have the insect tolerance of wild species, the first study of variation in the sequences of individual insect-sensitive genes across vulnerable cultivated germplasms and resistant wild cousins using multiomics techniques may be accomplished [120]. The resistance genes can be successfully used for gene editing after being validated against related insects. This presents chances for resistance development in the gene pool of cultivated crops to control insect pests [166]. It has been suggested that commercially valuable crops can produce insect-resistant phenotypes utilizing CRISPR-Cas gene-editing-based sequence variation by using either over-expression or silencing techniques. However, this has not yet been demonstrated.

4. Limitations and Future Perspectives

Like other biotechnological techniques, genome-editing techniques specifically modify a gene through cellular and in vitro mechanisms. In the course of evolution, genome modification is beyond our control. However, when the genome is altered experimentally, it may primarily be for the benefit of humans. Its application to crop improvement should, likewise, be limited to breeding objectives that are both absolutely important and challenging to achieve within the confines of the current heterogeneity. Like any modern technology, there are still a number of legal questions about gene editing that the scientific community needs to address. In order to fully utilize this innovation’s potential for the advancement of global agriculture and the eradication of neophobia in society, it is essential to adopt a realistic viewpoint that is supported by the legislative bodies that uphold scientific norms. The CRISPR-Cas-based deliberate dissemination of genetic components into wild species of insects that alter the population’s sex ratio or contribute to lethal mutations is a precise and environmentally sustainable method of battling pests. However, the emergence of insect resistance in response to a CRISPR-mediated gene drive could be a serious and ongoing problem at both the experimental and theoretical scales [167,168]. Multiplex gene editing, however, can overcome resistance [169]. Therefore, it is crucial to address insect resistance issues in order to reach an agreement on the ethics and science in favor of this technology.
Additionally, because engineered insect pests have the power to change the entire population or environment, the introduction of CRISPR-Cas-edited insects bearing gene drives into the ecosystem is linked to a number of biosafety concerns. Prior to their release, stringent risk assessments of non-target outcomes are also required. Unexpected post-release impacts on beneficial insects can have a negative influence on food chains and can alter the composition of communities [118]. Additionally, the disease can become worse due to the possibility of gene transfers between the target organisms and their non-target relatives. If the risks are appropriately managed in light of unanticipated environmental repercussions, gene-driven technology could prove effective in the targeted extermination of insect pests, insect vectors for viruses, andalien insect species. Utilizing the terminator genes that permit the programmed life of modified insects and using tagged insects to monitor gene flow seem to be crucial steps in the biosafe use of gene drives in the context of risk management. Additionally, another option for the management of invasive pests is the use of robotic equipment and artificial intelligence to physically eliminate individual pests [170]. Robotics may not be as effective, though, when dealing with tiny insects, uneven terrain, and hidden eggs.
Insect resistance to invasive pests has been successfully achieved via the CRISPR-Cas-based deletion of vulnerable genes. The fundamental problem associated with S gene deletions, which also add to the associated fitness penalty, is pleiotropic effects in the plant. However, it is possible to ensure insect resistance without affecting plant performance by altering the binding effector factor rather than the gene itself [171]. The CRISPR-Cas approach of creating insect resistance in crop species will, therefore, develop as a successful tool for supplying genetic traits in farmed varieties in a shorter amount of time. It is true that CRISPR-Cas-enabled genome-editing technology is a fast-evolving technique and, thus, the scope of its application in agriculture is expanding [172,173]. However, a thorough understanding of the gene and genome activities of the target species is required prior to its full adoption in the generation of insect pest resistance and plant protection. As Bt technology developed from recombinant DNA technology has revolutionized the management of insects in many economically important crops, including cotton, maize, soybean, and brinjal [174], the ease and multiplexing manner of CRISPR technology could also replace the currently used recombinant DNA technology for the insertion of R gene(s) in a faster manner.

5. Concluding Remarks

Despite being relatively young, the genome-editing techniques centered on CRISPR-Cas have already changed insects’ functional genomics. With CRISPR-Cas, we can now quickly alter, remove, and add DNA almost anywhere we want in any crop or insect species to make plants immune to insect pests. Therefore, this technology needs to be enhanced in order to produce crop plants that are resistant to insect pests. Sincere and proactive measures in this regard are required in addition to protecting our crops from the significant output losses brought on by insect pest infestation. However, the fate of genome-modified products with CRISPR in crop enhancement projects will ultimately be determined by the worldwide legislative authorities. For novel crop cultivars, either product- or process-based regulation is followed by regulatory systems. The scope of the regulations implemented on CRISPR-based crops will have an effect on the cost of their production and will also dictate the pace at which they will reach commercial industries. The set of product-based legislation for crops created using CRISPR-Cas genome editing could be classified similarly to products created by classical mutagenesis, eliminating them from the restrictions imposed on products made via genetic modification. This would surely have an impact on the hopeful public perception of this technology and would help the majority of nations to adopt it. Many countries have given the green pass to CRISPR-edited products that carry no transgene(s). It is expected that the CRISPR-Cas technology will lead to a new green revolution in agriculture if the timely deregulation of the adoption of CRISPR products and technological know-how is shared by an open scientific practice.

Author Contributions

Conceptualization, T.I.; formal analysis, T.T.M. and I.J.M.; investigation, T.T.M., M.C. and I.J.M.; resources, T.I.; data curation, T.I. and M.C.; writing—original draft preparation, T.T.M., M.C., A.A.M.K. and I.J.M.; writing—review and editing, T.I., M.C., M.S.U. and M.R.A.; visualization, T.I.; supervision, T.I. and M.R.A.; Funding acquisition, T.I. All authors have read and agreed to the published version of the manuscript.

Funding

This work was funded by the KrishiGobeshona Foundation (KGF) Bangladesh Project, project no. TF 50-C/17to T.I.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

All data used were included in the manuscript.

Acknowledgments

We are thankful to the KGF, Bangladesh, for funding this project.

Conflicts of Interest

The authors declared no conflict of interests.

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Stresses 02 00034 g001 550
Figure 1. Workflow for CRISPR-Cas9-based genome editing in insects and plants for insect resistance.
Figure 1. Workflow for CRISPR-Cas9-based genome editing in insects and plants for insect resistance.
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MDPI and ACS Style

Moon, T.T.; Maliha, I.J.; Khan, A.A.M.; Chakraborty, M.; Uddin, M.S.; Amin, M.R.; Islam, T. CRISPR-Cas Genome Editing for Insect Pest Stress Management in Crop Plants. Stresses 2022, 2, 493-514. https://doi.org/10.3390/stresses2040034

AMA Style

Moon TT, Maliha IJ, Khan AAM, Chakraborty M, Uddin MS, Amin MR, Islam T. CRISPR-Cas Genome Editing for Insect Pest Stress Management in Crop Plants. Stresses. 2022; 2(4):493-514. https://doi.org/10.3390/stresses2040034

Chicago/Turabian Style

Moon, Tasfia Tasnim, Ishrat Jahan Maliha, Abdullah Al Moin Khan, Moutoshi Chakraborty, Md Sharaf Uddin, Md Ruhul Amin, and Tofazzal Islam. 2022. "CRISPR-Cas Genome Editing for Insect Pest Stress Management in Crop Plants" Stresses 2, no. 4: 493-514. https://doi.org/10.3390/stresses2040034

APA Style

Moon, T. T., Maliha, I. J., Khan, A. A. M., Chakraborty, M., Uddin, M. S., Amin, M. R., & Islam, T. (2022). CRISPR-Cas Genome Editing for Insect Pest Stress Management in Crop Plants. Stresses, 2(4), 493-514. https://doi.org/10.3390/stresses2040034

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