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. 2026 Apr;189:104498. doi: 10.1016/j.ibmb.2026.104498

Knockout of nAChR subunits in spider mites and their phytoseiid predators confers spinosyn cross-resistance and reveals a conserved mode of action in mites

Antonio Mocchetti a, Pieter Steelant a, Mahboubeh Hosseinkhani b, Sander De Rouck a, Jahangir Khajehali a,c, Thomas Van Leeuwen a,⁎
PMCID: PMC12979025  PMID: 41577144

Abstract

Spinosyns are allosteric modulators of nicotinic acetylcholine receptors (nAChRs) which in insects specifically target subunit α6. However, their mode of action in mites and compatibility with phytoseiid predators remain unclear. We combined phylogenetics with CRISPR/Cas-based reverse genetics to test whether α6-like subunits mediate spinosyn toxicity in mites and to assess prospects for resistance breeding in phytoseiids. The phylogenetic analysis identified seven α and three β subunits in multiple phytoseiids and in Tetranychus urticae. A single phytoseiid subunit clustered within the insect α6/α7 clade, whereas T. urticae possessed three (Tuα5/α6/α7) without strict one-to-one insect orthology. Using SYNCAS maternal delivery of CRISPR RNPs, we disrupted the putative α6 ortholog in Amblyseius swirskii (Asα6) and each of the three α6/α7-clade genes in T. urticae. In A. swirskii, all survivors of a discriminating spinosad dose carried Asα6 indels, and three independently edited lines exhibited insensitivity to both spinosad and spinetoram (no significant mortality at 10 000 mg a.i./L), whereas the wild type showed LC50 = 163 mg/L (spinosad) and 54 mg/L (spinetoram). In T. urticae, Tuα6 knockouts conferred high cross-resistance to both compounds, while Tuα5 knockouts slightly increased susceptibility and Tuα7 knockouts produced modest resistance. Our data demonstrate that α6-mediated spinosyn action is conserved in mites, with α6 loss conferring strong cross-resistance in a key phytoseiid predator and in a model tetranychid. These findings enable marker-assisted editing/selection of spinosyn-resistant phytoseiid strains to improve pesticide–biocontrol compatibility and establish α6 as a practical universal marker gene for genome editing in acarine systems.

Keywords: Spinosad, Spinetoram, Insecticide resistance, CRISPR/Cas, Amblyseius swirskii, Tetranychus urticae, Microinjection

Graphical abstract

Image 1

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Highlights

  • •

    nAChRs subunit compositions differ between mites and insects.

  • •

    Knockout of α6 induce spinosyn resistance in A. swirskii and T. urticae.

  • •

    Knockout of Tuα5 and Tuα7 in T. urticae results in milder phenotypes.

  • •

    α6 is a practical universal marker gene for genome editing.

  • •

    Selective breeding on phytoseiid predators can develop spinosyn resistant strains.

1. Introduction

Integrated Pest Management (IPM) has gained worldwide recognition as an approach that suppresses pest populations while minimizing risks to human health and the environment. A cornerstone of IPM is augmentative biological control (ABC) which relies on the release of mass-reared biological control agents (BCAs) to regulate pest populations. Although pest management strategies based solely on BCAs have been proven successful in specific contexts (Bielza et al., 2020; van Lenteren et al., 2018), particularly in greenhouse systems, pesticide applications are often still required to keep pest damage below economic thresholds. Integrating BCAs and pesticides remains one of the main challenges of IPM programs, as BCAs can be severely affected by pesticides exposure, leading to reduced efficiency (Duso et al., 2020).

Phytoseiid mites (Acari: Phytoseiidae), represent over 60 % of the global market for arthropod natural enemies and are widely used to control insect and mite pests (Knapp et al., 2018; van Lenteren et al., 2018). Although phytoseiids are sometimes less affected by pesticide exposure than their prey, normally due to selective toxicity, they often still experience lethal or sublethal effects (Duso et al., 2020; İnak et al., 2025). Consequently, pesticide resistance has become a major trait of interest in BCA improvement programs, aimed to overcome current limitations of biological control (Bielza et al., 2020; Lirakis and Magalhães, 2019).

Understanding the molecular mechanisms underlying pesticide selectivity and resistance is crucial for advancing phytoseiid improvement through selective breeding or genetic engineering. CRISPR/Cas9 genome editing enables precise genetic modifications across a wide range of species. In arthropods, maternal delivery of Cas9 ribonucleoprotein has recently emerged as an effective alternative to traditional embryo injection (De Rouck et al., 2024; Li et al., 2021; Shirai et al., 2022). The SYNCAS method, originally developed for Tetranychus urticae, exploit the synergistic effect of saponins and branched amphiphilic peptide capsules (BAPC) to enhance delivery efficiency (De Rouck et al., 2024). This approach was the first to achieve successful gene knockouts in two phytoseiids species, Amblyseius swirskii and Phytoseiulus persimilis (Mocchetti et al., 2025a). By enabling targeted gene disruption and functional validation of candidate loci, CRISPR/Cas9 opens new opportunities for developing molecular markers and accelerating genetic improvement programs in biological control agents (Leung et al., 2020; Lirakis and Magalhães, 2019; Lv et al., 2025).

Spinosyns, such as spinosad and spinetoram act as allosteric modulators of nicotinic acetylcholine receptors (nAChRs) and are classified as group 5 insecticides according to IRAC MoA (Sparks et al., 2020). Spinosad, a fermentation product of the actinomycete Saccharopolyspora spinosa, consists of a mixture of spinosyn A and D. Spinetoram, a semi-synthetic derivative of the naturally occurring spinosyn J and L, exhibits enhanced residual activity and a broader pest spectrum (Galm and Sparks, 2016; Sparks et al., 2021). nAChRs belong to the Cys-loop ligand-gated ion channels family and are pentameric complexes that create a central ion pore, with diverse subunit combinations producing distinct receptor subtypes in arthropods (Breer and Sattelle, 1987; Le Novère et al., 2002). Spinosyns resistance has been extensively documented in insects and can arise from both metabolic detoxification and alteration of nAChR α6 subunit (Sparks et al., 2025). In particular, amino acid substitutions (e.g., G275E) or complete loss-of-function of α6 confer very high resistance levels in multiple insect species to both spinosad and spinetoram (Baxter et al., 2010; Mocchetti et al., 2025b; Puinean et al., 2013; Wan et al., 2018).

However, information on the effects of spinosyns and the underlying resistance mechanisms in mites remains limited. Some studies have reported off-target effects on phytoseiids and toxicity in Tetranychus urticae, showing that both compounds can induce lethal and sublethal effects in several species (Busuulwa et al., 2024; Kim et al., 2018; Schmidt-Jeffris et al., 2021; van Leeuwen et al., 2005), whereas recently, knockout of a putative nAChR α6 subunit in Neoseiulus californicus led to increased resistance to spinetoram while maintaining susceptibility to spinosad, suggesting that in this species spinosad mode of action might differ from insects.

This study aims to apply a reverse-genetic approach based on CRISPR/Cas-mediated gene knockout to target putative α6 subunits in mite species. Findings from A. swirskii, the most economically important phytoseiid, may provide guidelines for future genetic improvement programs aimed at developing pesticide-resistant strains of phytoseiid mites. In parallel, the use of the model mite T. urticae reinforces conclusions about the mode of action of spinosyns in mites.

2. Material and method

2.1. Mite rearing

A commercial strain of A. swirskii (Biobest, Belgium) and the German Susceptible Strain (GSS) of T. urticae (Stumpf et al., 2001) were used as reference populations. Both A. swirskii and T. urticae lines were reared on Phaseolus vulgaris cv. “Prelude” in mite-proof cages at room temperature with 16:8 light:dark photoperiod. Typha pollen (Nutrimite™, Biobest) was provided as food source for A. swirskii and occasionally supplemented with T. urticae to stimulate eggs production. After injections, mites were reared on detached bean leaves on wet cotton in plastic containers in climate chambers at 26 °C, 60 % relative humidity (RH) and 16:8 light:dark photoperiod.

2.2. Toxicity bioassays

Dose-response assays were performed by directly spraying on mites with a Cornelis spray tower (Van Leeuwen et al., 2004). Briefly, 20–30 mites were placed on leaf disks surrounded by a water barrier and sprayed with at least five concentrations of Spinosad (Conserve Pro™, Corteva Agriscience) or spinetoram (Exalt™, Corteva Agriscience). Every concentration was tested in at least four replicates. Mortality was scored 24 h after treatment for A. swirskii and 72 h after treatment for T. urticae by gently poking the mites with a small brush. These time points were selected based on preliminary assays indicating that more accurate LC50 estimates were obtained for T. urticae when mortality was scored after 72 h, whereas 24 h was the best for A. swirskii. Mites unable to walk three times their body length were considered dead. When 10 000 mg active ingredient (a.i.)/L did not cause 50 % mortality, higher concentrations were not tested to avoid potential interference from other components in the commercial formulations, which are not intended for use at such high doses.

2.3. Identification of nAChR subunits and phylogenetic analysis

Sequences of D. melanogaster were retrieved from FlyBase, (https://flybase.org/). Manually curated sequences from Apis mellifera, Lepeophtheirus salmonis, Frankliniella occidentalis, Varroa destructor, Ixodes ricinus and T. urticae were obtained from different publications (Dermauw et al., 2012; Jones and Sattelle, 2006; Rispe et al., 2022; Rufener et al., 2020; Wan et al., 2018). Sequences from Galendromus occidentalis and Neoseiulus californicus were obtained from the NCBI reference assemblies (GCF_000255335.2 and GCA_028455905.1) using subunits of D. melanogaster and T. urticae as queries for tblastn. Sequences from A. andersoni and P. persimilis were manually annotated and curated from in-house available unpublished genomes, while A. swirskii sequences were annotated from a published transcriptome (Paspati et al., 2022). However, as one of the nAChR subunit was missing in this transcriptome, specific primers to amplify the complete coding sequence were designed on conserved regions across phytoseiid orthologs. RNA was extracted from 100 adult females of A. swirskii with the RNeasy mini kit (Qiagen). Next, 2 μg of RNA were used for first strand cDNA synthesis with the Maxima First Strand cDNA synthesis kit for RT-PCR (Fermentas Life Sciences). PCR was conducted using As_full_fwd and As_full_rvs (Table S1) on cDNA with GoTaq G2 DNA polymerase (Promega, Madison, WI, USA) starting with a 3 min denaturation step at 94 °C followed by 38 cycles of 1 min 45 s at 94 °C, 30 s at 53 °C and 2 min at 72 °C. A final elongation step at 72 °C was held for 5 min. PCR product quality was evaluated by running a 2 % agarose gel at 100 V for 30 min, purified using the E.Z.N.A.® Cycle Pure Kit (Omega Bio-Tek) and Sanger sequenced at LGC Genomics (Berlin, Germany). The complete coding sequence is reported in File S1.

Maximum-likelihood phylogenetic inference was used because overall protein similarity metrics alone did not provide sufficient resolution to reliably assign orthology among nAChR subunits. Protein sequences were aligned using MAFFT v7.520 using “auto” option and including NtR of D. melanogaster, closely related to nAChR subunits, as outgroup. Maximum-likelihood (ML) phylogenetic analysis was performed using IQtree version 2.4.0 (Minh et al., 2020) with the integrated ModelFinder function and ultrafast bootstrap (UFBoot) set at 10 000 replicates (Hoang et al., 2018). Conventional insect nomenclature was used to name phytoseiids subunits based on the clustering within the tree (Mitchell et al., 2022). Subunits lacking two adjacent cysteines in loop C were annotated as β subunits. The tree was rooted using MEGA11 and customized with Iroki (Moore et al., 2020). Protein sequences of all subunits used to build the tree are reported in File S2.

2.4. sgRNAs design

Guide sequences were designed with the online tool CRISPOR (Concordet and Haeussler, 2018). Targets were selected in regions upstream or within trans-membrane (TM) domains that are not known to undergo alternative splicing in insects and mites. Guide sequences were designed selecting the protospacer adjacent motif (PAM) NGG, recognized by Cas9 nuclease from Streptococcus pyogenes. A guide sequence adjacent to the TTT(A/G/C) PAM site, which is recognized by Cas12, was selected to target α5 in T. urticae due to the presence of predicted off-targets for guide sequences adjacent to the standard NGG PAM sites. An overview of the guide sequences used in this study is provided in Table S1.

To verify the target sequences in the strains of A. swirskii and T. urticae, DNA was extracted from at least 50 mites using the DNeasy Blood & Tissue Kit (Qiagen) followed by PCR of the target regions. Amplifications were performed with the primers listed in Table S1 using GoTaq G2 DNA polymerase starting with a 3 min denaturation step at 94 °C followed by 38 cycles of 1 min at 94 °C, 30 s at 53 °C and 1 min at 72 °C. A final elongation step at 72 °C was held for 5 min. PCR products were purified and sequenced as described above.

2.5. CRISPR/Cas9 formulation and injection mix

The SYNCAS formulation was utilized for all injections (De Rouck et al., 2024; Mocchetti et al., 2025a). Recombinant S. pyogenes Cas9 protein (Alt-R® S.p. Cas9 Nuclease V3) and recombinant Lachnospiraceae bacterium Cas12a protein (Alt-R™ L.b. Cas12a (Cpf1) Ultra) were purchased from Integrated DNA Technologies (IDT, Leuven, Belgium) and concentrated to 50 μg/μL using an Amicon® Ultra Centrifugal Filter (30 kDa cut-off). sgRNAs (Alt-R™ CRISPR/Cas9 sgRNA, IDT) and crRNA (Alt-R™ L.b. Cas12a crRNA, IDT) were dissolved in TE buffer to reach a final concentration of 10 μg/μL and 4 μg/μL, respectively. BAPC (Phoreus Biotech) and saponins (Saponin from Quillaja sp., Sigma-Aldrich #S4521) were dissolved in nuclease-free water to a final concentration of 10 μg/μL and 2.6 μg/μL respectively.

Details about mixture preparation are described in De Rouck et al. (2024). Briefly, 1.5 μL of Cas9 (or Cas12) and 3 μL of sgRNA (or crRNA) were incubated at room temperature for 10 min. Then 0.5 μL of BAPC, and 0.2 μL of saponin were included in the mixture that was subsequently incubated in ice for 30 min. Finally, the mixture was centrifuged at 20 000 g for 5 min at 4 °C and kept on ice until injection.

2.6. Injections, screening of mutants and generation of stable lines

Injections in both A. swirskii and T. urticae were performed on unsynchronized fertilized females, following previously described protocols (De Rouck et al., 2024; Mocchetti et al., 2025a). For A. swirskii, injections targeting the putative α6 were conducted in three independent replicates to assess editing efficiency. For T. urticae, a single injection was performed for each target gene where at least 200 females were used. Injected G-1 mothers were transferred to a new leaf disk after 24 h and finally discarded after 48 h.

In A. swirskii, G0 offspring were reared to adulthood and subsequently sprayed with 1000 mg/L spinosad, previously assessed as discriminating dose able to kill 100 % A. swirskii individuals. Mites surviving after 24 h were considered mutant. Surviving males were crossed with virgin wild-type (WT) females, whereas surviving females were already fertilized by G0 males as fertilization is needed for egg laying. After five days, DNA was extracted from surviving mites by crushing single individuals in 20 μl of STE buffer (100 mM NaCl, 10 mM Tris-HCl, 1 mM EDTA, pH 8) supplemented with 2 μL of proteinase K (10 mg/mL). Crude extracts were incubated at 55 °C for 30 min followed by 10 min at 99 °C. Samples were stored at −20 °C until PCR was conducted. DNA extractions and PCRs were also performed on a subset of dead mites. Editing efficiencies were calculated separately for offspring produced within 24 and 48 h after injection. Stable lines were then generated through subsequent sibling crosses. When possible (i.e., when mutations were expected to be homozygous), a selection step was conducted with spinosad to identify edited mites (see Results).

In T. urticae, 60 G0 males were each paired with one deutonymph or teliochrysalis (i.e., virgin) G0 female, all originated from eggs laid 24–48 h after injection. After four to five days of oviposition, DNA was extracted from the individual mites used for the crosses as described above. PCRs, purification and Sanger sequencing were performed as described above. Based on the genotype of the crossed mites, subsequent sibling or backcross mating were executed.

3. Results

3.1. Phylogenetic analysis of nAChR subunits and identification of α6 in A. swirskii and T. urticae

Annotation of nAChR subunits in phytoseiid species revealed the presence of seven α subunits and three β subunits, all conserved among the species analyzed. The same number of subunits was identified in T. urticae (Dermauw et al., 2012), however, phylogenetic analysis showed that a strict one-to-one orthology between phytoseiid and T. urticae subunits was not present (Fig. 1). Phylogenetic reconstruction also indicated that many mite subunits lack clear insect orthologs, with the exception of β1, which displayed unambiguous orthology with insects’ β1, and α2 and α8, although their clustering is not supported by high UFBoot values. Interestingly, a duplication of α2 was detected in all phytoseiid species examined but was absent in closely related species such as V. destructor and I. ricinus.

Fig. 1.

Fig. 1

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Maximum-likelihood phylogenetic tree of nAChR subunits of arthropod species. Included species are D. melanogaster (Dm), A. mellifera (Am), F. occidentalis (Fo), L. salmonis (Ls), V. destructor (Vd), I. ricinus (Ir), A. swirskii (As), A. andersoni (An), P. persimilis (Pp), G. occidentalis (Go) and T. urticae (Tu). Only UFBoot values > 95 are shown. Green dots indicate phytoseiids subunits whereas red dots indicate T. urticae subunits. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

The insect α6/α7 cluster, which also includes α5 of D. melanogaster, formed a well-supported clade with a single ortholog from Parasitiformes mites (including all phytoseiids species), and three orthologs from T. urticae. Based on the tree topology, mite subunits clustered more closely with insects α7, although node support was relatively low (UFBoot <95). Consequently, phylogenetic analysis was not sufficient to identify a clear α6 ortholog in mites. Nevertheless, because only a single phytoseiid subunit clustered within the α6/α7 clade, it was selected as the target for genome editing in A. swirskii. In T. urticae, the subunits annotated as α5, α6 and α7 formed a distinct subclade and their precise orthology with insect α6 remained uncertain. Also in this case it was not possible to determine which subunit was the functional ortholog of α6. Therefore, all three subunits were targeted with CRISPR/Cas9.

Other mite subunits also lacked clear orthologs. For α1, two distinct clades were observed: one comprising Parasitiformes and another comprising Pancrustacea. The T. urticae α1 sequence clustered separately. No mite orthologs were found for insects’ α3 and α4. Several α and β subunits were highly divergent overall.

3.2. Knockout of putative α6 subunits in A. swirskii and T. urticae and generation of stable lines

Both Asα6 and Tuα5, Tuα6, Tuα7 were targeted using the SYNCAS formulation in two reference strains. In A. swirskii, three independent injection replicates were performed to obtain a more accurate estimate of knockout efficiency, while in T. urticae, where the method is established with high efficacy, a single replicate injection was carried out for each gene.

Between 200 and 400 fertilized A. swirskii females were injected per replicate (Table 1). G0 male and female offspring were subsequently screened using a discriminating dose of spinosad. All survived mites, six in total, exhibited CRISPR-induced mutations, including both in-frame and out-of-frame events (chromatograms available in File S3.1). Editing efficiency attested around 3 % of offspring originated after 24 h. However, in replicate 2 the only mutant was identified in offspring originated within 24 h. No CRISPR events were detected in any of the 97 dead mites analyzed (File S3.2). This is surprising, as heterozygous females were expected among the dead mites (spinosad resistance mediated by α6 knockout is recessive in insects).

Table 1.

Editing efficiencies (percentage) obtained in A. swirskii targeting Asα6 with SYNCAS.

Rep. Injected mothers Alive after 24h Alive after 48h Offspring 0–24h Edited 0–24h Efficiency 0–24h Offspring 24–48h Edited 24–48h Efficiency 24–48h
1 201 118 109 50 0 0 86 3 3.49
2 386 232 223 27 1 3.7 68 0 0
3 400 192 183 32 0 0 66 2 3.03
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Three mutant mites were used to establish independent edited lines (Fig. 2, File S3.3): α6-KO-A carried an 11-nt deletion with a single nucleotide substitution, α6-KO-B carried a 7-nt insertion, and α6-KO-D carried a 6-nt deletion (Fig. 3).

Fig. 2.

Fig. 2

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Schematic representation of the successive sibling crosses used to generate three independent stable knockout lines in A. swirskii. The figure shows only the cross combinations relevant for establishing stable lines. Light-colored mites represent individuals from the parental WT strain. Red and green rectangles indicate knockout events. When possible (i.e., when mutations were expected to be homozygous), offspring carrying knockout alleles were selected through spinosad applications. In α6-KO-D, only male offspring were obtained after the first cross; therefore, an additional cross with a WT female was required. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Fig. 3.

Fig. 3

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Gene model of nAChR subunits targeted with CRISPR/Cas. Asα6 (A), Tuα5 (B), Tuα6 (C) and Tuα7 (D) and CRISPR-induced mutations in knockout stable lines. Introns indicated with dotted lines are not to scale. The four transmembrane domains (TM) are indicated with rectangles: red for TM1, blue for TM2, magenta for TM3 and green for TM4. Expected cleavage sites of Cas9 and Cas12 (Tuα5) are indicated with scissors. Letters in red indicate insertions or nucleotide substitutions. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

In T. urticae, only one injection was performed for each target gene, followed by random crosses between G0 males and females and subsequent marker-assisted backcrossing to establish stable lines (Fig. S1). Editing efficiencies, calculated based only on genotyped mites were 18.9 % for α5, 42.4 % for α6, and 18.1 % for α7. Three stable lines were obtained for each target gene (Fig. 3). All Tuα6 knockout lines presented the same, independently induced mutation (4-bp deletion). Independent onset of identical recurrent mutation was already observed in T. urticae and it is likely mediated by the microhomology-mediated end joining repair pathway which is believed to be dominant in this species.

3.3. Spinosyn toxicity

Dose-response curves were obtained for both A. swirskii and T. urticae parental and knockout lines (Table 2). In A. swirskii, the parental strain exhibited LC50 values of 163 mg/L for spinosad and 54 mg/L for spinetoram. Knockout of α6 conferred complete insensitivity to both spinosyns. LC50 values could not be determined, as even the highest concentration tested (10 000 mg/L) caused no significant mortality compared to the water control. Furthermore, oviposition seemed not to be affected by treatments.

Table 2.

LC50 values for A. swirskii BSS parental and knockout lines.

Line n LC50 (95 % CI) Slope ± SE χ2 (df) RR
Spinosad
 WT 776 163 (125–205) 2.35 ± 0.17 111.56 (44) 1
α6-KO-A 80 >10 000 / / >61
α6-KO-B 80 >10 000 / / >61
α6-KO-D
 80
 >10 000
 /
 /
 >61
Spinetoram WT 915 54.1 (43.7–65.1) 2.20 ± 0.14 133.14 (48) 1
α6-KO-A 80 >10 000 / / >185
α6-KO-B 80 >10 000 / / >185
α6-KO-D 80 >10 000 / / >185
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In T. urticae, α6 knockout also resulted in increased resistance to both spinosad and spinetoram (Table 3). Interestingly, while spinetoram was more toxic than spinosad in the parental strain, the opposite pattern was observed in knockout lines, where LC50 values for spinetoram exceeded those for spinosad. This result contrasts with all published studies in insects, where spinetoram consistently shows greater toxicity than spinosad even in α6 loss-of-function mutants.

Table 3.

LC50 values for T. urticae parental and knockout lines.

Chemical Line n LC50 (95 % CI) Slope ± SE χ2 (df) RR
Spinosad WT 478 40.6 (25.8–52.7) 2.06 ± 0.27 28.68 (18) 1
α5-KO-28 420 29.7 (23.4–41.7) 2.90 ± 0.35 60.65 (18) 0.73 (0.55–0.96)
α5-KO-38 427 22.4 (18.2–26.8) 4.66 ± 0.40 78.58 (18) 0.55 (0.43–0.71)
α5-KO-47
 504
 19.7 (16.3–22.8)
 4.40 ± 0.39
 57.94 (18)
 0.48 (0.37–0.62)
α6-KO-13 615 3720 (2440–6490) 1.64 ± 0.17 80.56 (18) 91.5 (67.1–125)
α6-KO-23 633 3430 (2900–4040) 3.97 ± 0.27 95.98 (22) 84.5 (65.6–109)
α6-KO-45
 641
 3990 (2820–6370)
 1.47 ± 0.15
 55.50 (18)
 98.2 (71.6–135)
α7-KO-10 436 66.9 (53.4–79.7) 4.12 ± 0.39 58.82 (18) 1.64 (1.27–2.14)
α7-KO-33 569 125 (98.4–169) 2.16 ± 0.21 60.99 (18) 3.07 (2.33–4.04)
α7-KO-47
 602
 90.5 (73.0–109)
 3.73 ± 0.32
 77.14 (18)
 2.23 (1.72–2.88)
Spinetoram WT
 688
 10.1 (7.70–12.7)
 2.49 ± 0.18
 88.23 (27)
 1
α5-KO-28 438 3.60 (2.93–4.17) 3.96 ± 0.39 44.51 (18) 0.35 (0.30–0.42)
α5-KO-38 479 3.95 (2.10–4.81) 4.17 ± 0.39 81.24 (18) 0.39 (0.33–0.46)
α5-KO-47
 464
 4.55 (3.99–5.13)
 4.91 ± 0.39
 47.54 (18)
 0.45 (0.39–0.52)
α6-KO-13 466 5890 (4490–8390) 2.70 ± 0.45 38.33 (18) 581 (470–718)
α6-KO-23 463 4740 (3540–5720) 3.70 ± 0.67 32.44 (18) 467 (385–567)
α6-KO-45
 536
 4610 (2680–11000)
 1.52 ± 0.23
 85.69 (20)
 454 (345–599)
α7-KO-10 494 12.7 (9.51–16.9) 3.82 ± 0.29 126.05 (18) 1.24 (1.05–1.46)
α7-KO-33 531 19.0 (15.2–23.2) 2.85 ± 0.23 40.15 (18) 1.87 (1.56–2.25)
α7-KO-47 523 20.6 (14.2–30.1) 1.87 ± 0.16 80.75 (18) 2.04 (1.66–2.49)
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By contrast, α5 and α7 knockout mutants exhibited less pronounced phenotypic changes (Table 3). Nonetheless, the response was consistent across independent lines: all α5 KO lines showed a slight increase in susceptibility to spinosyns, whereas all α7 knockout lines displayed a modest increase in resistance.

4. Discussion

Phytoseiid mites are widely used in IPM programs to control major arthropod pests. However, the implementation of genetic improvement programs is urgently needed to overcome current limitations of biological control, which are often caused by abiotic and biotic stresses affecting the performance of BCAs (Bielza et al., 2020; Lirakis and Magalhães, 2019). Several studies have shown that spinosyns can severely reduce the fitness of phytoseiid mites, potentially compromising their effectiveness as BCAs (Busuulwa et al., 2024; Kim et al., 2018; Schmidt-Jeffris et al., 2021). The development of spinosyn-resistant strains of phytoseiids would help to overcome this limitation, which is particularly relevant in organic agriculture, where both phytoseiid mites and spinosad are commonly used to control pest populations. While the molecular determinants of spinosyn toxicity and resistance are well characterized in insects, their mode of action in mites remains largely unexplored (Perry et al., 2025; Sparks et al., 2025). Number and composition of nAChR subunits can vary substantially across taxa, making it difficult to predict the role of specific subunits in pesticide resistance. Recently α6 knockout in the phytoseiid N. californicus suggested that an alternative mode of action of spinosyns was possible in mite species (Lv et al., 2025).

To investigate this, α and β subunits in several phytoseiid species were annotated and subsequently a comprehensive ML phylogenetic analysis was performed to determine relationships with other arthropod subunits, including other mite species as T. urticae, V. destructor and I. ricinus (Fig. 1). Only a few clear insect orthologs were identified in mites. Notably, no unambiguous α6 ortholog was found in any mite species; instead, putative α6 mite subunits formed a clade together with insect α6 and α7 (and α5 of Diptera). In D. melanogaster, knockout of α6 alone confers complete spinosyn resistance, whereas knockout of the closely related α5 or α7 subunits does not (Lu et al., 2022; Perry et al., 2021). The function of mite subunits within this α6/α7-related clade therefore remained uncertain and required functional validation.

Phytoseiid mites and other Parasitiformes contained only a single α6/α7-like subunit, whereas T. urticae possessed three which clustered separately (Fig. 1). To assess their functional role, Asα6 and Tuα5, Tuα6, Tuα7 were all disrupted using the SYNCAS CRISPR/Cas system.

In A. swirskii, G1 offspring were exposed to a discriminating dose of spinosad that caused 100 % mortality in WT individuals. All surviving mites carried mutations around the Cas9 cleavage site, confirming that disruption of Asα6 confers spinosad resistance (File S3.1). The clear phenotypic distinction between resistant and WT individuals allowed Asα6 to serve as a reliable marker gene for estimating editing efficiency, something not previously feasible in phytoseiids. Editing efficiency varied among the three replicates targeting α6 but appeared consistent (3–4 %) with previous results based solely on genotyping (Mocchetti et al., 2025a). Surprisingly, in replicate two, only a single edited mite was detected in the 0–24 h offspring, and none were found in the 24–48 h group (Table 1).

Interestingly, no heterozygous individuals carrying a single edited α6 allele were detected among dead A. swirskii mites. In the study of De Rouck et al. (2024) it was demonstrated that when fertilized females were injected, more than 2 events could be found in the G0 daughters indicating Cas9 activity occurred after the first mitotic division which occurs only several hours after oviposition. This indicates the Cas9 remains active for a considerable time after being taken up by the eggs. Hence, in predatory mites the Cas9 enzyme might have sufficient time to cleave both alleles in the eggs where Cas9 is successfully delivered, making the occurrence of heterozygous or chimeric mutants rare. However, given that only 97 dead individuals were sequenced, heterozygotes could still be present among untested specimens. Although the efficiency of SYNCAS in phytoseiids does not yet reach that of T. urticae, this study demonstrates that the method is already suitable for routine KO applications. Furthermore, the identification of α6 as an appropriate marker gene will facilitate optimizing the method for phytoseiid species.

To assess the strength of the resistant phenotype induced by Asα6 disruption, dose-response curves were obtained for parental and knockout lines (Table 2). In the WT strain, spinetoram was more toxic than spinosad, consistent with observations in other arthropods, likely due to its higher affinity for α6 and potential secondary targets (Sparks et al., 2008). In contrast, all three knockout lines were completely insensitive to both compounds. LC50 values could not be determined as even at 10 000 mg/L, the highest concentration tested, no significant mortality was observed. Interestingly, α6-KO-D, which carried an in-frame mutation causing the loss of two amino acids and substitution of one in proximity of TM1 (Fig. 3), displayed the same resistance phenotype as other knockouts, suggesting that even minor alterations in these regions can abolish receptor function.

These results in A. swirskii (and T. urticae, discussed below) suggest a conserved mode of action for spinosyns in insects and mites. However, they contrast with recent findings in N. californicus, where α6 knockout conferred resistance to spinetoram but not to spinosad (Lv et al., 2025). The peculiar phenotype observed in N. californicus warrants further investigation, as the available information does not clearly explain its case. Early studies on α6 knockout in insects hypothesized that truncated/partial subunits might retain some function, for example by sequestering spinosyns in resistant individuals (Baxter et al., 2010; Wan et al., 2018), potentially accounting for differences between compounds. However, complete deletion of α6 in D. melanogaster resulted in full resistance (Zimmer et al., 2016), suggesting that loss of the canonical target site alone explains resistance. Furthermore, in the study of Lv and colleagues, large deletions were generated using two sgRNAs, resulting in loss of the region spanning from exon 2 to exon 10, likely eliminating all spinosyns binding sites. Therefore, the persistence of susceptibility to spinosad when α6 is absent implies the existence of a secondary target site. Yet, subunit compositions and amino acid sequences seem highly conserved among phytoseiids (Fig. 1). Although unlikely, minor amino acid differences in alternative subunits might have generated a secondary target site of spinosad in N. californicus that is absent in other mites or insects, where α6 loss consistently confers cross-resistance. Complete susceptibility of the Ncα6 knockout line indicates that spinosad could target this hypothetical secondary target with the same efficiency as α6. Alternatively, the line might have an Ncα6 duplication that is not disrupted and repeated selection with spinetoram might have fixed a spinetoram-specific (metabolic) resistance gene that is physically near the Ncα6 targeted region and hence co-segregates in F2 genetic linkage screens. These results put forward the need to create replicate lines with different independent events.

Although results reported in the present study are consistent with observations in insects, where knockout of α6 always translated in high resistance levels for both compounds (Mocchetti et al., 2025b; Shi et al., 2022; Zimmer et al., 2016; Zuo et al., 2020), they remain surprising as no close orthologs of Asα6 were identified. This poses the question of which subunits can compensate for α6 loss. It is well known that nAChR subunits maintain a certain degree of redundancy, as demonstrated by non-lethal knockout mutants (Korona et al., 2022; Lu et al., 2022; Perry et al., 2021), although redundancy has to be expected among subunits part of the same subclass (Le Novère et al., 2002). In D. melanogaster, α5 and α7 are closely related to α6 and likely form heteromeric nAChRs, however, their single and combined knockouts never induced spinosad resistance, indicating that α6 could still form spinosyn responsive receptors in the absence of the two subunits (Lansdell et al., 2012; Lu et al., 2022; Watson et al., 2010). Even though functional homomeric α6 receptors have been obtained ex vivo (Hawkins et al., 2022; Khan et al., 2025; Turberg et al., 2021), it remains unclear whether such assemblies exist in vivo (Korona et al., 2022). Notably, the effect of triple α5/α6/α7 knockouts in D. melanogaster has never been tested. Such experiments could clarify whether functional redundancy is confined to these subunits or extends to more distantly related ones. Here, α6 knockout in a mite lacking closely related orthologs suggests that α6 function may be partially or fully compensated by more divergent subunits, a hypothesis that warrants further investigation across arthropods.

To extend these findings, putative α6 subunits were also targeted in T. urticae, a model mite species. Here, the focus lied on generating stable knockout lines, as editing efficiency in this species is well established (De Rouck et al., 2024). Notably, the targeting of Tuα5 with SYNCAS was performed using Cas12 instead of Cas9 that was consistently used in past experiments. This decision was driven by the possible off-target predicted for Cas9 sgRNAs. Comparable editing efficiencies between Cas12 and Cas9 suggest that Cas12 is a suitable alternative endonuclease for future SYNCAS applications. Three independent lines were produced for each gene to determine knockout phenotypes (Fig. 3).

Strikingly, all Tuα6 knockout lines displayed high resistance to both compounds. However, the LC50 values for spinetoram were consistently higher than those for spinosad, resulting in resistance ratios (RR) of 454–580 compared to 84–98 for spinosad. This pattern appears to be unique to mites, as in all insect species where α6 knockouts have been generated, spinetoram has consistently shown greater toxicity than spinosad (e.g., Mocchetti et al., 2025b; Shi et al., 2022; Zuo et al., 2020). These findings suggest that spinetoram may interact with an additional neuronal target in insects that is absent in Acari. Consequently, the higher toxicity of spinetoram observed in WT mites may simply reflect its superior stability and stronger affinity for Tuα6. Moreover, the relatively higher toxicity of spinosad in Tuα6 knockouts (i.e., when the primary molecular target site is absent) could indicate greater metabolic stability or partial interaction with alternative low-affinity targets. A recent patent application indicated α5 of a tick species as affected by spinosad binding (Turberg et al., 2021). T. urticae (but not phytoseiid mites) present a close ortholog of ticks α5, here named Tuα3 (Fig. 1). Whether this could act as secondary for spinosad and spinetoram would require further investigations.

In Tuα5 and Tuα7 lines phenotypic changes were less pronounced, but consistent among replicates and across compounds. An increase in susceptibility (0.4–0.7 fold) was observed for Tuα5 knockout. This might be explained by the substitution in nAChRs of the nonfunctional Tuα5 subunits with Tuα6, increasing the absolute number of nAChRs including Tuα6 in the nervous system of T. urticae and consequently the toxicity of spinosyns. Conversely, Tuα7 knockout caused a minor resistance increase (RR 1.24–3.07), suggesting that Tuα7 may serve as a secondary, low-affinity spinosyn target. This hypothesis aligns with the naturally present substitution in Tuα7 of the canonical glycine at position 275 with alanine (known to confer resistance in D. melanogaster), likely reducing spinosyn affinity (Scott et al., 2023). Additionally, Tuα7 shows low expression across all life stages (Dermauw et al., 2012), implying a limited physiological role. Its close genomic linkage (7 kbp) to Tuα6 suggests a recent duplication event in T. urticae.

Further studies are required to gain a deeper understanding of subunit composition of mite nAChRs. Ex vivo heterologous expression of non-hybrid nAChRs would be crucial for elucidating how different subunits assemble and for investigating their roles as pesticide targets. Although this has long been considered challenging in arthropods (Ihara et al., 2020; Rufener et al., 2020), recent evidence of successful heterologous expression of homomeric α6 nAChRs from two tick species, without the need of ancillary proteins, suggests that expression of mite subunits might be more straightforward (Cartereau et al., 2025; Khan et al., 2025; Turberg et al., 2021). Several other aspects remain to be elucidated, including the intrinsic structural features that make α6 the only nAChR subunit efficiently targeted by spinosyns, the potential presence and role of alternative splicing, and the fitness costs associated with α6 disruption in mite species.

Overall, these results indicate that the molecular mode of action of spinosyns is conserved between insects and most mites, confirming α6 as principal molecular target. The discovery that α6 knockout in A. swirskii confers a high level of resistance to spinosyns established the foundation for developing spinosyn-resistant phytoseiid strains for IPM. However, due to current limitations on the use of genetically edited animals in fields and greenhouses (e.g., in Europe), the direct release of such strains would not be permitted. Alternatively, naturally occurring α6 knockout mutations could be identified in wild populations or induced through mutagenesis, and subsequently introgressed into commercial strains via marker-assisted selective breeding. Such strains could be deployed alongside conventional chemical controls to enhance biological control efficacy, taking advantage of the environmentally friendly profile of spinosad which is approved for organic agriculture. In addition, these resistant predatory mites might slow down the development of resistant pest populations in fields that are repeatedly sprayed with spinosyns as individuals surviving pesticide treatment could still be eliminated via predation and the required number of pesticide treatments would be expected to be lower, reducing selection pressure. Finally, the clear phenotypic marker conferred by α6 disruption, and the conserved nature of this mechanism, highlights its potential as a universal marker gene for genome editing studies in arthropods susceptible to spinosyns.

CRediT authorship contribution statement

Antonio Mocchetti: Writing – original draft, Visualization, Validation, Methodology, Investigation, Formal analysis, Data curation. Pieter Steelant: Writing – review & editing, Investigation, Formal analysis, Data curation. Mahboubeh Hosseinkhani: Investigation, Formal analysis, Data curation. Sander De Rouck: Writing – review & editing, Investigation, Formal analysis, Data curation. Jahangir Khajehali: Investigation, Formal analysis, Data curation. Thomas Van Leeuwen: Writing – review & editing, Funding acquisition, Conceptualization.

Declaration of generative AI in scientific writing

During the preparation of this work the consensus search engine was used to find relevant literature: https://consensus.app/search/

Declaration of interests

The authors declare that there are no competing interests.

Acknowledgements

This work was supported by ERC Proof of Concept [grant 101123162-CRISPART to T.V.L.], Research Foundation Flanders (FWO) [G0A1525N and G017923N] and by a bilateral research cooperation between Flanders (FWO) – Vietnam (NAFOSTED) [grant G0E1221N to T.V.L]. Sander De Rouck is a post- doctoral fellow of FWO (grant 1237426N).

Footnotes

Appendix A

Supplementary data to this article can be found online at https://doi.org/10.1016/j.ibmb.2026.104498.

Contributor Information

Antonio Mocchetti, Email: antonio.mocchetti@ugent.be.

Pieter Steelant, Email: pieter.steelant@ugent.be.

Mahboubeh Hosseinkhani, Email: mhosseinkhani@ut.ac.ir.

Sander De Rouck, Email: sander.derouck@ugent.be.

Jahangir Khajehali, Email: khajeali@cc.iut.ac.ir.

Thomas Van Leeuwen, Email: thomas.vanleeuwen@ugent.be.

Appendix A. Supplementary data

The following are the Supplementary data to this article:

Multimedia component 1
mmc1.docx (198.4KB, docx)
Multimedia component 2
mmc2.zip (768B, zip)
Multimedia component 3
mmc3.zip (21.6KB, zip)
Multimedia component 4
mmc4.zip (14.1MB, zip)

Data availability

Data will be made available on request.

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Associated Data

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Supplementary Materials

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mmc2.zip (768B, zip)
Multimedia component 3
mmc3.zip (21.6KB, zip)
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Data Availability Statement

Data will be made available on request.

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