Dominant point mutation in a tetraspanin geneassociated with field-evolved resistance ofcotton bollworm to transgenic Bt cottonLin Jina,1, Jing Wanga,1, Fang Guana, Jianpeng Zhanga, Shan Yua, Shaoyan Liua, Yuanyuan Xuea, Lingli Lia, Shuwen Wua,Xingliang Wanga, Yihua Yanga, Heba Abdelgaffarb, Juan Luis Jurat-Fuentesb, Bruce E. Tabashnikc, and Yidong Wua,2

aCollege of Plant Protection, Nanjing Agricultural University, Nanjing, Jiangsu, China; bDepartment of Entomology and Plant Pathology, University ofTennessee, Knoxville, TN 37996; and cDepartment of Entomology, University of Arizona, Tucson, AZ 85721

Edited by May R. Berenbaum, University of Illinois at Urbana–Champaign, Urbana, IL, and approved October 5, 2018 (received for review July 15, 2018)

Extensive planting of crops genetically engineered to produce in-secticidal proteins from the bacterium Bacillus thuringiensis (Bt) hassuppressed some major pests, reduced insecticide sprays, enhancedpest control by natural enemies, and increased grower profits. How-ever, rapid evolution of resistance in pests is reducing these benefits.Better understanding of the genetic basis of resistance to Bt crops isurgently needed to monitor, delay, and counter pest resistance. Wediscovered that a point mutation in a previously unknown tetraspa-nin gene in the cotton bollworm (Helicoverpa armigera), a devastat-ing global pest, confers dominant resistance to Cry1Ac, the sole Btprotein produced by transgenic cotton planted in China. We foundthe mutation using a genome-wide association study, followed byfine-scale genetic mapping and DNA sequence comparisons betweenresistant and susceptible strains. CRISPR/Cas9 knockout of the tetra-spanin gene restored susceptibility to a resistant strain, whereasinserting the mutation conferred 125-fold resistance in a susceptiblestrain. DNA screening of moths captured from 23 field sites in sixprovinces of northern China revealed a 100-fold increase in the fre-quency of this mutation, from 0.001 in 2006 to 0.10 in 2016. Thecorrespondence between the observed trajectory of the mutationand the trajectory predicted from simulation modeling shows thatthe dominance of the mutation accelerated adaptation. Proactiveidentification and tracking of the tetraspanin mutation demonstratethe potential for genomic analysis, gene editing, and molecular mon-itoring to improve management of resistance.

evolution | resistance management | genetically modified | dominance |sustainability

Genetically engineered crops that produce insecticidal proteinsfrom the bacterium Bacillus thuringiensis (Bt) have been

planted globally on a cumulative total of over 930 million hectaressince 1996 (1). The benefits of these transgenic Bt crops includepest suppression, reduced insecticide use, enhanced biologicalcontrol, and increased farmer profits (2–7). However, increasinglyrapid evolution of resistance to Bt crops by pests has eroded thesebenefits (8–10). Better understanding of the genetic basis of re-sistance to Bt crops is urgently needed to monitor, delay, andcounter pest resistance.The most widely adopted strategy for delaying pest resistance to

Bt crops entails “refuges” of non-Bt host plants that enable survivalof susceptible insects to mate with resistant insects (8). Refuges areespecially effective for delaying resistance that is inherited as arecessive trait, because the matings between rare homozygous re-sistant insects and relatively abundant homozygous susceptible in-sects from refuges produce heterozygous progeny that are killed bythe Bt crop (11). Because such recessive resistance can be sup-pressed more readily, nonrecessive resistance is more likely toevolve in the field (8). Nonetheless, most research has focused onrecessive resistance conferred by mutations that disrupt binding ofBt toxins in the larval midgut to receptors such as cadherin andATP-binding cassette transporter proteins (12–17), whereas little isknown about the genetic basis of dominant resistance. Moreover,

previous efforts to achieve proactive molecular monitoring of Btresistance have had limited success because the mutations thatincrease markedly in the field are usually identified after resistancehas caused severe control failures (13, 14, 16).Here, we report the discovery and proactive monitoring of a

point mutation in a tetraspanin gene that confers dominant re-sistance to Bt toxin Cry1Ac in the cotton bollworm, Helicoverpaarmigera. This lepidopteran is one of the world’s most devastatingcrop pests and has recently invaded the Americas (18, 19). Weanalyzed H. armigera from northern China, where Bt cotton pro-ducing Cry1Ac has been planted by millions of smallholderfarmers since 1997 (20, 21). The percentage of H. armigera larvaeresistant to Cry1Ac increased significantly there, from 0.93% in2010 to 5.5% in 2013 (21). This has been categorized as an “earlywarning of resistance,” because the percentage of resistant indi-viduals did not exceed 50% and reduced efficacy of Bt cotton inthe field was not reported (8, 21).Previous work showed the resistance to Cry1Ac in the AY2

strain of H. armigera from northern China was autosomal,dominant, and 1,200-fold relative to the susceptible strain SCD,

Significance

Crops genetically engineered to produce insecticidal proteinsfrom the bacterium Bacillus thuringiensis (Bt) kill some majorpests and reduce use of insecticide sprays. However, evolutionof pest resistance to Bt proteins decreases these benefits. Betterunderstanding of the genetic basis of resistance to Bt crops isurgently needed to address this problem. We discovered that apoint mutation in the cotton bollworm, one of the world’s mostvoracious pests, confers dominantly inherited resistance to theBt protein produced by transgenic cotton grown in China. Thismutation increased 100-fold in frequency from 2006 to 2016 inChina. Proactive tracking of this mutation may improve man-agement of resistance and enhance sustainability of Bt cottonfor millions of smallholder farmers in China.

Author contributions: Y.Y. and Y.W. designed research; L.J., J.W., F.G., J.Z., S.Y., S.L., Y.X.,L.L., H.A., and J.L.J.-F. performed research; S.W., X.W., and Y.Y. contributed new reagents/analytic tools; L.J., J.W., F.G., J.Z., B.E.T., and Y.W. analyzed data; B.E.T. conducted mod-eling; and L.J., J.W., B.E.T., and Y.W. wrote the paper.

Conflict of interest statement: B.E.T. is coauthor of a patent on modified Bacillus thur-ingiensis toxins, “Suppression of Resistance in Insects to Bacillus thuringiensis Cry Toxins,Using Toxins That Do Not Require the Cadherin Receptor” (patent nos. CA2690188A1,CN101730712A, EP2184293A2, EP2184293A4, EP2184293B1, WO2008150150A2, andWO2008150150A3). Amvac, Bayer CropScience, Dow AgroSciences, DuPont Pioneer, Mon-santo, and Syngenta did not provide funding to support this work, but may be affectedfinancially by publication of this paper and have funded other work by B.E.T.

This article is a PNAS Direct Submission.

Published under the PNAS license.1L.J. and J.W. contributed equally to this work.2To whom correspondence should be addressed. Email: wyd@njau.edu.cn.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1812138115/-/DCSupplemental.

Published online October 31, 2018.

11760–11765 | PNAS | November 13, 2018 | vol. 115 | no. 46 www.pnas.org/cgi/doi/10.1073/pnas.1812138115

Dow

nloa

ded

by g

uest

on

Oct

ober

27,

202

0

based on the concentration of Cry1Ac killing 50% of larvae (LC50)(20). The dominance parameter, h, which varies from 0 for com-pletely recessive to 1 for completely dominant, was previouslyreported for AY2 as 1.0 based on larval survival at the diagnostictoxin concentration (1 μg of Cry1Ac per square centimeter of diet)(20). In this study, we calculated h as 0.79 (SI Appendix, Supple-mentary Methods), based on a more comprehensive approach us-ing data from previous bioassays on Bt cotton for survival fromneonate to adult, sex ratio, and fertile eggs per female (21).Here, a genome-wide association study (GWAS) with 48 re-

sistant larvae and 48 susceptible larvae from a mass backcrossexperiment using AY2 and SCD revealed a highly significant as-sociation with resistance to Cry1Ac for four of 2,097 single-nucleotide polymorphisms (SNPs), all from 10.48 to 11.15 Mbp onchromosome 10 (Fig. 1A and SI Appendix, Table S1). Next, weconducted fine-scale mapping using seven other SNPs from 10.58to 11.01 Mbp on chromosome 10 and 363 larvae that were derivedfrom three single-pair backcross families and survived exposure tothe diagnostic toxin concentration (SI Appendix, Fig. S1B). Theresults demonstrate a highly significant association between re-sistance and each of these seven SNPs (P < 10−60 for each SNP;Fig. 1B and SI Appendix, Table S2).We narrowed our search to the 17 of 21 genes between 10.62

and 10.87 Mbp on chromosome 10 that are expressed in themidgut of final instar larvae (SI Appendix, Table S3). We designedspecific primers to amplify by PCR the complete ORF of these 17genes from the cDNA of the resistant AY2 strain and the sus-ceptible SCD strain. Comparison of the predicted amino acidsequences between the two strains identified a single amino acidsubstitution (L31S) in a tetraspanin gene of AY2, and no other

mutations in the 17 genes that differed consistently betweenstrains. We named this geneHaTSPAN1 (GenBank accession nos.MH514007 for SCD and MH514008 for AY2).Tetraspanins are a family of proteins that are important in cell

migration, signal transduction, and intracellular trafficking, as wellas infection by diverse pathogens, including bacteria (22, 23). Thefull-length transcript of HaTSPAN1 from SCD encodes a proteinof 304 amino acids with 63.4% identity to the 23-kDa integralmembrane protein of Bombyx mori (GenBank accession no.XP_004933861). The predicted structure of the HaTSPAN1protein includes characteristic features of tetraspanins: fourtransmembrane segments (TM1–TM4), one small and one largeextracellular loop (EC1 and EC2, respectively), short intracellularamino and carboxyl tails, and the signature CCG motif (Fig. 1C).The L31S substitution occurs in the third amino acid of TM1 (Fig.

1 C and D) and is encoded by DNA near the 3′-end of exon 1 (Fig.2A). Comparison of cDNA sequences between AY2 and SCD indi-cates the L31S substitution is encoded by a single nucleotide sub-stitution: T92C. Analysis of the 363 backcross progeny that survivedexposure to the diagnostic concentration in the fine-scale mappingexperiment (SI Appendix, Fig. S1B) confirmed a strong associationbetween T92C and resistance (345 heterozygous for T92C, 18 ho-mozygous for wild type; χ2 = 298.6, df = 1, P = 5 × 10−66). Survival ofthe 18 larvae that lacked the T92C mutation apparently was con-ferred by other mutations, nongenetic factors, or both.The leucine at position 31 predicted in wild-type HaTSPAN1

is conserved in this protein and homologous proteins from sus-ceptible strains of 10 species representing seven families ofLepidoptera (SI Appendix, Fig. S2) and in the SCD-r1 strain ofH. armigera, which has recessive resistance to Cry1Ac conferred

Fig. 1. Identification of the mutation in HaTSPAN1associated with dominant resistance to Cry1Ac in H.armigera. (A) GWAS based on 2,097 SNPs in the AY2strain. The horizontal line shows the threshold forsignificant association (P < 5 × 10−8). The two mostsignificant points have the same probability (10−11)and nearly identical positions in chromosome 10 (SIAppendix, Table S1); they are shifted slightly apart sothat both are visible. (B) SNP markers and HaTSPAN1locus on HaChr10. (C) HaTSPAN1 protein with fourtransmembrane domains (TM1–TM4), two extracellu-lar loops (EC1 and EC2), and the L31S mutation inTM1. (D) Amino acid sequence alignment of the TM1domain of HaTSPAN1 in five strains of H. armigera[two with the wild-type (WT) sequence (the suscepti-ble SCD strain and the SCD-r1 strain with recessivecadherin-based resistance) and three with the L31Smutation (MUT; AY2, SCD423, and QX7 with dominantresistance)] and homologous genes from nine otherlepidopteran species from seven families: Helicoverpazea (Noctuidae); Papilio machaon, Papilio polytes,Papilio xuthus (Papilionidae); Danaus plexippus(Nymphalidae); Pectinophora gossypiella (Gelechiidae);Amyelois transitella (Pyralidae); Plutella xylostella(Plutellidae); and Bombyx mori (Bombycidae).

Jin et al. PNAS | November 13, 2018 | vol. 115 | no. 46 | 11761

AGRICU

LTURA

LSC

IENCE

S

Dow

nloa

ded

by g

uest

on

Oct

ober

27,

202

0

by a cadherin mutation (Fig. 1D). Conversely, the T92C muta-tion yielding the L31S substitution occurs in AY2 and two otherstrains of H. armigera from northern China (SCD423 and QX7)that have dominant resistance to Cry1Ac (Fig. 1D). We detectedthe T92C mutation in all 36 larvae tested from SCD423 (23homozygotes and 13 heterozygotes) and in all 28 larvae testedfrom QX7 (23 homozygotes and five heterozygotes).The results from GWAS, fine-scale mapping, and sequence

comparisons reported above show a strong association betweenT92C and resistance to Cry1Ac in several strains of H. armigera.However, like nearly all related previous work, they do not dem-onstrate causation. Here, we used gene editing with CRISPR/Cas9to test the hypothesis that the T92C mutation in HaTSPAN1causes resistance to Cry1Ac. First, we created knockout strainAY2-KO in which a 4-bp insertion in exon 4 (Fig. 2A) introduceda premature stop codon expected to produce a truncated, inactiveHaTSPAN1 protein. Of 835 AY2 eggs injected with a single-guideRNA (sgRNA1; SI Appendix, Table S4) and Cas9 protein, 78%hatched and 66% of the neonates developed to adults (G0). Re-ciprocal mass crosses between moths from G0 and SCD producedthe next generation (G1). Among the 96 G1 pupae genotyped withnondestructive exuviate-based PCR, 74% were heterozygous forindel mutations in HaTSPAN1. From these heterozygotes, fivefemales and four males harboring the 4-bp insertion were pooledto generate the G2 moths. Of the G2 moths genotyped by exuviate-based PCR, 27 of 122 (22%) were homozygous for the 4-bp in-sertion. We pooled these 27 moths to establish the homozygousknockout strain AY2-KO. AY2-KO was as susceptible to Cry1Acas the susceptible SCD strain (Fig. 3 and SI Appendix, Table S5),indicating the knockout of HaTSPAN1 eliminated resistance.Next, we used CRISPR/Cas9 to introduce the T92C mutation

into SCD and create knock-in strain SCD-KI. Of 1,800 SCD eggsinjected with sgRNA2, Cas9, and a single-stranded oligodeox-ynucleotide, 26 hatched, and 17 of the neonates developed toadults. Reciprocal mass crosses between these G0 adults and SCDproduced G1 larvae that had a survival rate of 1.9% (95 of 4,896)at the diagnostic concentration of Cry1Ac. The 95 survivors were

reared to adults and mass-crossed to generate G2 larvae that had asurvival rate of 25% (581 of 2,304) at the diagnostic concentration.The 288 largest of these survivors (≥10 mg) were reared to pu-pation and nondestructively genotyped: 157 were homozygousfor the T92C mutation, and the rest were heterozygous. The157 homozygous mutant moths were mass-crossed to establishthe SCD-KI strain. Relative to SCD, SCD-KI had 125-foldresistance to Cry1Ac (Fig. 3 and SI Appendix, Table S5), dem-onstrating that the T92C mutation can cause resistance to Cry1Ac.Survival at the diagnostic concentration was 81% (n = 48) for

Fig. 2. CRISPR/Cas9 editing of HaTSPAN1. (A) Redarrows show positions of mutations introduced withCRISPR/Cas9. (B) Portion of the wild-type (WT) se-quence and a single-stranded oligodeoxynucleotide(ssODN) used as donor DNA to introduce four single-base substitutions (red letters in boxes) in exon 1. TheT92C mutation yields the L31S amino acid substitu-tion. The other three substitutions are synonymousand serve as markers, including one in the PAM se-quence (highlighted in green) that also preventsdouble cutting. The sgRNA target sequence is high-lighted in yellow. (C) Chromatograms of direct se-quencing of PCR products for determining genotype.

Fig. 3. Responses to Cry1Ac by five strains of H. armigera. AY2, resistant;AY2-KO, AY2 with knockout of HaTSPAN1; SCD, susceptible; SCD423, re-sistance introgressed into SCD; SCD-KI, SCD with knock-in of the T92C muta-tion in HaTSPAN1. The resistance ratio is the LC50 for a strain divided by theLC50 for the SCD strain. The resistance ratio did not differ significantly betweenSCD (1) and AY2-KO (0.9). Details are provided in SI Appendix, Table S5.

11762 | www.pnas.org/cgi/doi/10.1073/pnas.1812138115 Jin et al.

Dow

nloa

ded

by g

uest

on

Oct

ober

27,

202

0

SCD-KI, 45% (n = 288) for the first generation (F1) progeny fromcrosses between SCD-KI and SCD, and 0% for SCD (n = 48),which yields h = 0.56 for SCD-KI.Relative to SCD-KI, resistance to Cry1Ac was higher and more

dominant in AY2. At the diagnostic concentration, AY2 survivalwas 96% and h was 1.0 (20). Also, the LC50 of Cry1Ac was ninefoldgreater for AY2 than SCD-KI (Fig. 3 and SI Appendix, Table S5).These results suggest that one or more factors other than the T92Cmutation boosted dominance and resistance in AY2 relative toSCD-KI. However, AY2-KO had no resistance to Cry1Ac, in-dicating that CRISPR/Cas9 introduction of the mutation yielding atruncated HaTSPAN1 protein completely restored susceptibility.Considered together, these results imply that one or more factorsin AY2 other than the T92C mutation can interact with that mu-tation to increase dominance and resistance, but such factorsconfer little or no resistance in the absence of the T92C mutation.One such factor could be the 2.7-fold increased transcription ofHaTSPAN1 in AY2 relative to SCD (P < 10−19; SI Appendix, TableS3), which might increase resistance in combination with T92C butis not expected to confer resistance without T92C.The AY2 strain had 10-fold cross-resistance to Bt toxin

Cry2Ab (SI Appendix, Table S5), which does not share bindingsites with Cry1Ac in H. armigera (24). AY2-KO was not signifi-cantly cross-resistant to Cry2Ab, indicating that the knockout ofHaTSPAN1 restored susceptibility to Cry2Ab. However, SCD-KIalso was not significantly cross-resistant to Cry2Ab. These resultssuggest that one or more factors in AY2 interact with the T92Cmutation to cause cross-resistance to Cry2Ab, but they do notconfer significant cross-resistance without the T92C mutation.Unlike the markedly reduced binding of Cry1Ac in the SCD-r1

strain of H. armigera and many other strains with recessive re-sistance to Bt toxins (12), substantially reduced binding of Cry1Acwas not associated with the dominant resistance of AY2 (SI Ap-pendix, Fig. S3). This supports the idea that the T92C mutationinvolves a gain of function, rather than a loss of function, andsuggests interference with other steps in the toxic pathway (25).To test the hypothesis that the T92C mutation in HaTSPAN1

contributes to resistance to Bt cotton in the field, we used twomethods to track its frequency in ethanol-preserved H. armigeramoths captured from 2006 to 2016 at 23 sites in six provinces ofnorthern China (SI Appendix, Fig. S4). In the first of two screeningmethods, we tested moths in pools to enable efficient screening,using amplicon sequencing. We tested a total of 5,996 mothscaptured in seven different years (2006, 2010, and 2012–2016) byscreening pools, with a mean of 102 moths per pool (SE = 8). Inthe second method, to check the effectiveness of screening pools,we used the more rigorous and labor-intensive approach of testingmoths individually. The 2,259 moths tested individually were asubset of the moths captured during 2006, 2010, 2013, and 2016.Consistent with the hypothesis that the T92C mutation con-

tributes to resistance to Bt cotton in the field, both methodsrevealed a significant increase in the frequency of the T92C mu-tation from 2006 to 2016 (Mann–Whitney U test, P < 0.005 foreach method). Based on screening pooled moths, the frequency ofT92C increased 100-fold, from 0.001 [95% confidence interval(CI): 0.0–0.003] in 2006 to 0.10 (95% CI: 0.05–0.15) in 2016 (Fig. 4and SI Appendix, Table S6). Based on screening moths individually,the frequency increased from 0.0 in 2006 (none detected, n = 454moths) to 0.093 (95% CI: 0.06–0.12) in 2016 (SI Appendix, TableS7). For the 4 y that both methods were applied, the results arecorrelated between the two methods (r = 0.98, df = 2, P = 0.019; SIAppendix, Fig. S5), and no significant difference occurred betweenthem in any year (Mann–Whitney U test, P > 0.6 for each year).Based on pooled moths, the percentage of populations evaluatedthat had at least one T92C mutation increased from 12% (one ofeight) in 2006, to 50% (three of six) in 2010, to 88% (seven ofeight) in 2016 (SI Appendix, Table S6).To evaluate the importance of the dominance of the resistance

conferred by the T92C mutation, we compared the observed tra-jectory of this mutation with the trajectories predicted by simula-tion modeling under three different assumptions: the empirically

determined dominant resistance (h = 0.79) and hypothetical sce-narios with either recessive inheritance of resistance (h = 0) orintermediate dominance (h = 0.4) (Fig. 4 and SI Appendix, TableS8). With dominant resistance, the predicted trajectory corre-sponded well with the observed trajectory, but with h = 0 or 0.4, thepredicted frequency increased much more slowly than the observedfrequency (Fig. 4). A sensitivity analysis varying assumptions aboutdominance and the fitness cost associated with resistance showsthese results are robust (SI Appendix, Fig. S6), confirming thecritical role of the dominance of the T92C mutation in acceleratingthe evolution of resistance. Moreover, among survivors of exposureto the diagnostic concentration of Cry1Ac, the percentage of indi-viduals that were either heterozygous or homozygous for T92Cincreased from 44% in 2011 to 70% in 2016 (SI Appendix, Fig. S7).These data demonstrate a faster increase in frequency for T92Crelative to the other mutations contributing to field-evolved re-sistance, including the recessive cadherin mutations (26).Although the T92C frequency in H. armigera increased 100-fold

to 0.10 in 2016, reduced efficacy of Bt cotton against this pest hasnot been reported yet in northern China. However, analysis ofthe observed T92C frequency data from 2012 to 2016 indicatesthat if the current trajectory continues, the T92C frequency innorthern China will exceed 0.50 by 2023 (linear regression of log-transformed data: r2 = 0.91, df = 3, P = 0.01). The tracking ofT92C reported here may spur changes to delay resistance, whiletracking it in the future could help to determine if such changesare effective. Options for slowing resistance include switching totransgenic cotton that augments Cry1Ac with one or two unrelatedBt toxins (Cry2Ab and Vip3Aa) or RNA interference (27, 28),increasing the abundance of non-Bt host plant refuges, and in-tegrating diverse tactics for pest control (8, 29).As far as we know, previous work has not reported interactions

between Bt toxins and tetraspanins. Web of Science searchesidentified over 24,000 papers on Bt and over 5,000 on tetra-spanins, but none on both. More work is needed to determinethe function of wild-type HaTSPAN1, how the T92C mutationconfers resistance, and if mutations in tetraspanin genes conferdominant resistance to Bt toxins in other insects. With resistancean urgent global threat to food security and human health (30),the results here demonstrate the potential for genomic analysis,

Fig. 4. Observed versus predicted frequency of the T92C mutation in H.armigera in northern China. The predicted values are from simulations of apopulation genetic model with dominance (h) set to hypothetical values of0 (recessive) or 0.4 (intermediate dominance), or the empirically determinedvalue of 0.79 (dominant). The observed values are means (with 95% CIs)based on DNA screening of pooled moths. For all simulations, the observedfrequency for 2006 of 0.001 (95% CI: 0.0–0.003) was used as the initial fre-quency (i.e., for 2006; SI Appendix, Table S8).

Jin et al. PNAS | November 13, 2018 | vol. 115 | no. 46 | 11763

AGRICU

LTURA

LSC

IENCE

S

Dow

nloa

ded

by g

uest

on

Oct

ober

27,

202

0

gene editing, and molecular monitoring to facilitate moresustainable pest control.

Materials and MethodsInsect Strains. We used seven strains of H. armigera: resistant strains AY2,QX7, SCD423, SCD-r1, and SCD-KI and susceptible strains SCD and AY2-KO.All strains were maintained as described previously, with larvae reared on anartificial diet and adults provided with 10% sugar solution (31). The twoprimary strains of H. armigera analyzed here are the previously describedresistant strain AY2 and susceptible strain SCD. AY2 was started in 2011 bypairing a susceptible SCD female with a resistant male from Anyang in theHenan province of northern China (20). The descendants from this single-pair mating were selected with increasing concentrations of Cry1Ac (20).SCD was started with insects from Côte D’Ivoire (Ivory Coast) over 30 y agoand has been maintained in the laboratory without exposure to Bt toxins orother insecticides (32). In diet overlay bioassays with Cry1Ac, the LC50 was1,200-fold higher for AY2 than SCD and resistance at the diagnostic con-centration was dominant (h = 1) (20).

We also analyzed three other previously described resistant strains: QX7 andSCD423 with dominant resistance and SCD-r1 with recessive resistance. In parallelwith AY2, QX7 was started in 2011 by pairing a susceptible SCD female with aresistant male from Qiuxian in the Hebei province of northern China, followed byselection of subsequent generations with Cry1Ac (20). Resistance to Cry1Ac in QX7was 460-fold relative to SCD and dominant (h = 1) (20). SCD423 originated fromcrosses between SCD and resistant strain AY423, whichwas established from an F2screen of a field population from Anyang in 2009 (26). Resistance to Cry1Ac inAY423 was 660-fold relative to SCD and dominant (h = 0.64) (26). The resistanceto Cry1Ac in AY423 was introgressed into SCD by four crosses with SCD, followedby selection with Cry1Ac (26). Although the resistance to Cry1Ac was similar inSCD423 and AY423, SCD423 shares ca. 94% of its genetic background with SCD.The previously described SCD-r1 strain had 500-fold resistance to Cry1Ac caused byintrogressing the recessive r1 cadherin allele (h = 0) into SCD (32).

Here, we created the susceptible AY2-KO strain and the resistant SCD-KIstrain by CRISPR/Cas9-mediated editing. AY2-KOwas created by knocking outHaTSPAN1 from AY2, and SCD-KI was created by knocking in the T92Cmutation into SCD.

Bt Toxins and Bioassays. For bioassays, we used activated Cry1Ac toxin pur-chased from Marianne Pusztai-Carey (Case Western Reserve University,Cleveland, OH) and Cry2Ab protoxin generously provided by the Institute ofPlant Protection, Chinese Academy of Agricultural Sciences, Beijing, China. Weused diet overlay bioassays as described previously (20). Toxin stock suspensionswere diluted with a 0.01 M (pH 7.4) phosphate buffer solution (PBS). Liquidartificial diet (900 μL) was dispensed into each well (surface area = 2 cm2) of a24-well plate. After the diet cooled and solidified, 100 μL of PBS containing thedesired concentration of Cry1Ac or Cry2Ab was applied evenly to the dietsurface in each well and allowed to dry. A single larva was placed in each well.Larvae were kept at 26 °C (±1°), 60% (±10%) relative humidity, and 16 h light:8 h dark. We followed established protocols for each toxin (31). For Cry1Ac, wetested second instars that had been starved for 4 h and recorded mortalityafter 5 d. For Cry2Ab, we tested unfed neonates (24 h old) and recordedmortality after 7 d. At the end of the bioassays, larvae were scored as dead ifthey died or weighed less than 5 mg.

GWAS of Resistance to Cry1Ac in AY2.Backcross. We conducted a mass cross with 30 females from the resistant AY2strain and 30 males from the susceptible SCD strain (SI Appendix, Fig. S1A).Crossing over occurs only in male Lepidoptera (33, 34), and resistance toCry1Ac in AY2 is dominant (20). Accordingly, we generated backcrossprogeny (BC1) by allowing mating between 30 females from SCD and 30 F1males from AY2 × SCD.Phenotyping by bioassay. To distinguish between susceptible and resistantindividuals, 240 larvae from BC1 were reared on diet treated with 0.25 μg ofCry1Ac per square centimeter of diet and 240 were reared on diet with 10-fold that concentration (SI Appendix, Fig. S1A). After 5 d, 53 live larvaeexposed to the lower concentration that weighed <5 mg were scored assusceptible (BC1-S), while 97 live larvae exposed to the higher concentrationthat weighed >10 mg were scored as resistant (BC1-R). Larvae from eachgroup were reared to fifth instars on untreated diet for DNA extraction.DNA extraction. We dissected 48 midguts from fifth instars from each group,washed them in ice-cold 0.15 M NaCl solution, and extracted DNA from eachmidgut by the phenol/chloroform method. Each sample was digested in-dividually in 500 μL of DNA lysis buffer [100mM Tris, 50mM EDTA, 200mMNaCl, 1% SDS (pH 8.0)] for 10 min at 56 °C. We purified DNA by extraction

three times with phenol/chloroform (1:1 vol/vol), and then once with phenol.After centrifugation at 12,000 × g for 10min, the supernatant was collectedand DNA was precipitated using 50 μL of NH4COOH (pH 7.5) and 800 μL of100% ethanol. The DNA pellet was washed with 500 μL of 70% ethanol,resuspended in 80 μL of TE buffer [1mM EDTA, 10mM Tris (pH 8.0)], andstored at −20 °C.Genotyping by sequencing. DNA was digested by the Pstl restriction enzyme,which recognizes a 6-bp sequence (5′-CTGCAG-3′). Genomic DNA libraries wereprepared for genotyping by sequencing by ligating the digested DNA tounique nucleotide barcode adapters for each individual and a commonadapter for all individuals. Next, we pooled the DNA samples and conductedPCR amplification (35). Single-end sequencing (85-bp reads) of the 96-plex li-brary per flow cell channel was performed on a Genome Analyzer II (Illumina,Inc.). We performed an initial quality check using FastQC (www.bioinformatics.babraham.ac.uk/projects/fastqc/). We used Process_Radtags built-in Stacks (36)for demultiplexing and cleaning of sequencing reads. Wemapped high-qualityreads to the H. armigera genome (37) using the Burrows–Wheeler Aligner (38)and called SNPs using the Genome Analysis Toolkit (39). We assigned 2,097segregating SNPs to the 31 chromosomes of H. armigera.Association between resistance and SNPs. We performed GWAS using PLINKwith stringent quality-control filters (40). We used individuals and SNPs thatmet three criteria: individuals with 50% of all called SNPs, loci that werepresent in 50% of individuals, and a minor allele frequency of >0.03. Weanalyzed 2,097 informative SNPs from 93 individuals to compare allele fre-quencies between BC1-R and BC1-S using PLINK. We obtained P values fromFisher’s exact test and produced a Manhattan plot of the results using thesoftware package “qqman” (41).

Fine-Scale Mapping of Resistance. For fine-scale mapping of resistance withinH. armigera chromosome 10 (HaChr10), we conducted a second backcross(BC2) (SI Appendix, Fig. S1B). As with BC1 described above, we started with amass cross between 30 females from AY2 and 30 males from SCD (SI Ap-pendix, Fig. S1B). Backcross families (BC2) were obtained by crossing F1(AY2 × SCD) males with SCD females in single pairs. The progeny from threesingle-pair BC2 families were allowed to feed on diet that had the diagnostictoxin concentration (1 μg of Cry1Ac per square centimeter of diet). Wenamed the group of 363 survivors BC2-R. Total genomic DNA was extractedindividually, as described above, from each of the 363 BC2-R survivors andthe parents of the three BC2 families.

Based on the genome sequence data of H. armigera (37), we designedspecific primers in the exons of functional genes spanning 10.58–11.01 Mbpon HaChr10. PCR amplification of the genomic DNA and sequencing wereperformed for each parent of the three BC2 families. As informative markers,we used each of the seven SNPs that were heterozygous in the father (F1 ofAY2 × SCD) and homozygous in the mother (SCD). We compared the ob-served frequency of each of these markers with the expected 1:1 frequencyif they segregated independent of resistance.

Larval Midgut RNA Abundance for 21 Genes from 10.62 to 10.87 Mbp onChromosome 10. We extracted total midgut RNA from final instars of SCDand AY2 using TRIzol (Invitrogen). Purified RNA was treated with DNase I toeliminate DNA contamination. From each strain, we obtained three replicateswith 30 midguts per replicate.

The mRNA was isolated from total RNA with oligo (dT) magnetic beads,and fragmented to generate reads that covered the entire length of thetranscripts using fragmentation buffer on a thermomixer. Subsequently,cDNA libraries (one library per sequencing sample) were constructed usingclassical Illumina protocols (42). Briefly, these cleaved mRNA fragments wereprimed randomly and subjected to the synthesis of the first-strand andsecond-strand cDNAs. Single-nucleotide A was added to obtain short frag-ments, the adapters were ligated to short fragments, and PCR was thenperformed. The quality of total RNA, mRNA, and cDNA libraries was checkedwith an Agilent 2100 Bioanalyzer instrument. The cDNA libraries were se-quenced on an Illumina HiSeq 2000 platform.

The raw reads were cleaned by omitting adapter sequences, reads withmore than 5%ambiguous bases N, and low-quality sequences withmore than15% of nucleotides with a Phred quality score of<20. The clean reads wereassembled de novo using Trinity software (43). We used SOAPaligner soft-ware for short oligonucleotide alignment to remove sequences that werenot covered by any sample reads. The assembled transcripts were processedthrough the TGICL Gene Indices clustering tools to eliminate sequence re-dundancy and further assemble sequences to generate effective unigenes(44). We assigned a gene ID to each of the 21 unigenes based on homologyto the genome database of H. armigera (37) identified with Blastx software.

11764 | www.pnas.org/cgi/doi/10.1073/pnas.1812138115 Jin et al.

Dow

nloa

ded

by g

uest

on

Oct

ober

27,

202

0

Sequencing Candidate Genes in the Region of Chromosome 10 Associated withResistance. Based on the gene sequences in the genome database of H.armigera (37), we designed specific primers in the 5′ and 3′ UTRs to amplify thecomplete ORF in AY2 and SCD of the 17 candidate genes that are expressed inthe midguts of fifth instars and occur from 10.62 to 10.87 Mbp on chromo-some 10. The PCR products were sequenced directly. The amino acid sequencewas predicted from the complete ORF sequence and aligned between AY2and SCD using DNAssist 2.2 software (https://dnassist.en.softonic.com/).

Predicted HaTSPAN1 Protein Structure. We used the amino acid sequence ofHaTSPAN1 to predict the protein’s structure, including TM helices usingTMHMM Server v2.0 (www.cbs.dtu.dk/services/TMHMM/). The conserveddomains were analyzed by Blastp to the Conserved Domain Database ofNCBI (https://www.ncbi.nlm.nih.gov/cdd).

Detection of the T92C Mutation of HaTSPAN1.We used the phenol/chloroformmethod described above to prepare genomic DNA from individual larvae(fourth or fifth instars) or moths from three strains (AY2, SCD423, and QX7)and from the 363 survivors of exposure to the diagnostic concentration in thefine-scale mapping experiment (SI Appendix, Fig. S1B). Specific primers TM1-F (forward) and TM1-R (reverse) (SI Appendix, Table S4) amplified a frag-ment of ∼500 bp containing exon 1 and partial intron 1 to detect the T92Cmutation of HaTSPAN1. The PCR mixture (20 μL) consisted of 10 μL of 2×PrimeSTAR Max Premix (TaKaRa), 1 μL of genomic DNA, 1 μL of 10 μM for-ward primer (TM1-F), 1 μL of 10 μM reverse primer (TM1-R), and 7 μL ofddH2O. The PCR conditions were as follows: 94 °C for 2 min followed by 35

cycles of 94 °C for 10 s, 55 °C for 30 s, and 72 °C for 1 min, followed by a finalextension at 72 °C for 10 min. The PCR products of the expected size weredirectly sequenced with the forward primer TM1-F by TSINGKE.

HaTSPAN1 Amino Acid Sequence Alignment in H. armigera and Nine OtherSpecies of Lepidoptera. The sequence alignment in Fig. 1D shows the re-gion of TM1 of the HaTSPAN1 protein containing the L31S mutation fromfive strains of H. armigera (mutant: AY2, QX7, and SCD423; wild type: SCDand SCD-r1) and homologs of HaTSPAN1 (with GenBank accession nos.MH514007 and MH514008) from Helicoverpa zea (NFMG01022088.1), Pap-ilio machaon (XP_014368154.1), Papilio polytes (XP_013141160.1), Papilioxuthus (XP_013182388.1), Danaus plexippus (EHJ70576.1), Pectinophoragossypiella (JAT80322.1), Amyelois transitella (XP_013190067.1), Plutellaxylostella (XP_011564311.1), and B. mori (XP_004933861).

Additional details of the methods for CRISPR/Cas9 editing of HaTSPAN1,binding of Cry1Ac, DNA screening of field-captured moths, and computersimulations are provided in SI Appendix, Supplementary Methods.

ACKNOWLEDGMENTS. We thank Y. Carrière, G. Davidowitz, A. Mathias,L. Matzkin, and S. Morin for suggestions that improved the paper. This workwas supported by the National Natural Science Foundation of China (Grant31530060), the Ministry of Agriculture and Rural Affairs of China (Grant2016ZX08012-004), the Agriculture and Food Research Initiative Program(Grant 2018-67013-27821), and the Biotechnology Risk Assessment ResearchGrant Program (Grant 2014-33522-22215) from the US Department of Agri-culture National Institute of Food and Agriculture.

1. James C (2017) Global Status of Commercialized Biotech/GM Crops in 2017: BiotechCrop Adoption Surges as Economic Benefits Accumulate in 22 Years (InternationalService for the Acquisition of Agri-biotech Applications, Ithaca, NY), ISAAA Brief No. 53.

2. Hutchison WD, et al. (2010) Areawide suppression of European corn borer with Btmaize reaps savings to non-Bt maize growers. Science 330:222–225.

3. Downes S, Walsh T, Tay WT (2016) Bt resistance in Australian insect pest species. CurrOpin Insect Sci 15:78–83.

4. Dively GP, et al. (2018) Regional pest suppression associated with widespread Btmaize adoption benefits vegetable growers. Proc Natl Acad Sci USA 115:3320–3325.

5. Lu Y, Wu K, Jiang Y, Guo Y, Desneux N (2012) Widespread adoption of Bt cotton andinsecticide decrease promotes biocontrol services. Nature 487:362–365.

6. National Academies of Sciences, Engineering, and Medicine (2016) GeneticallyEngineered Crops: Experiences and Prospects (National Academies Press, Wash-ington, DC).

7. Wu KM, Lu YH, Feng HQ, Jiang YY, Zhao JZ (2008) Suppression of cotton bollworm inmultiple crops in China in areas with Bt toxin-containing cotton. Science 321:1676–1678.

8. Tabashnik BE, Carrière Y (2017) Surge in insect resistance to transgenic crops andprospects for sustainability. Nat Biotechnol 35:926–935.

9. Gassmann AJ, et al. (2014) Field-evolved resistance by western corn rootworm tomultiple Bacillus thuringiensis toxins in transgenic maize. Proc Natl Acad Sci USA 111:5141–5146.

10. Tabashnik BE (2016) Tips for battling billion-dollar beetles. Science 354:552–553.11. Gould F (1998) Sustainability of transgenic insecticidal cultivars: Integrating pest ge-

netics and ecology. Annu Rev Entomol 43:701–726.12. Wu Y (2014) Detection and mechanisms of resistance evolved in insects to Cry toxins

from Bacillus thuringiensis. Adv Insect Physiol 47:297–342.13. Mathew LG, et al. (2018) ABC transporter mis-splicing associated with resistance to Bt

toxin Cry2Ab in laboratory- and field-selected pink bollworm. Sci Rep 8:13531.14. Banerjee R, et al. (2017) Mechanism and DNA-based detection of field-evolved re-

sistance to transgenic Bt corn in fall armyworm (Spodoptera frugiperda). Sci Rep 7:10877.

15. Walsh T, James B, Chakroun M, Ferré J, Downes S (2018) Isolating, characterising andidentifying a Cry1Ac resistance mutation in field populations of Helicoverpa puncti-gera. Sci Rep 8:2626.

16. Fabrick JA, et al. (2014) Alternative splicing and highly variable cadherin transcriptsassociated with field-evolved resistance of pink bollworm to Bt cotton in India. PLoSOne 9:e97900.

17. Peterson B, Bezuidenhout CC, Van den Berg J (2017) An overview of mechanisms ofCry toxin resistance in lepidopteran insects. J Econ Entomol 110:362–377.

18. Kriticos DJ, et al. (2015) The potential distribution of invading Helicoverpa armigerain North America: Is it just a matter of time? PLoS One 10:e0119618, and erratum(2015) 10:e0133224.

19. Anderson CJ, et al. (2018) Hybridization and gene flow in the mega-pest lineage ofmoth, Helicoverpa. Proc Natl Acad Sci USA 115:5034–5039.

20. Jin L, et al. (2013) Dominant resistance to Bt cotton and minor cross-resistance to Bttoxin Cry2Ab in cotton bollworm from China. Evol Appl 6:1222–1235.

21. Jin L, et al. (2015) Large-scale test of the natural refuge strategy for delaying insectresistance to transgenic Bt crops. Nat Biotechnol 33:169–174.

22. Monk PN, Partridge LJ (2012) Tetraspanins: Gateways for infection. Infect Disord DrugTargets 12:4–17.

23. Florin L, Lang T (2018) Tetraspanin assemblies in virus infection. Front Immunol 9:1140.

24. Hernández-Rodríguez CS, Van Vliet A, Bautsoens N, Van Rie J, Ferré J (2008) Specificbinding of Bacillus thuringiensis Cry2A insecticidal proteins to a common site in themidgut of Helicoverpa species. Appl Environ Microbiol 74:7654–7659.

25. Pardo-López L, Soberón M, Bravo A (2013) Bacillus thuringiensis insecticidal three-domain Cry toxins: Mode of action, insect resistance and consequences for cropprotection. FEMS Microbiol Rev 37:3–22.

26. Zhang H, et al. (2012) Diverse genetic basis of field-evolved resistance to Bt cotton incotton bollworm from China. Proc Natl Acad Sci USA 109:10275–10280.

27. Carrière Y, Fabrick JA, Tabashnik BE (2016) Can pyramids and seed mixtures delayresistance to Bt crops? Trends Biotechnol 34:291–302.

28. Ni M, et al. (2017) Next-generation transgenic cotton: Pyramiding RNAi and Btcounters insect resistance. Plant Biotechnol J 15:1204–1213.

29. Tabashnik BE, et al. (2010) Suppressing resistance to Bt cotton with sterile insect re-leases. Nat Biotechnol 28:1304–1307.

30. Gould F, Brown ZS, Kuzma J (2018) Wicked evolution: Can we address the sociobio-logical dilemma of pesticide resistance? Science 360:728–732.

31. Zhang H, et al. (2011) Early warning of cotton bollworm resistance associated withintensive planting of Bt cotton in China. PLoS One 6:e22874.

32. Yang YH, et al. (2009) Introgression of a disrupted cadherin gene enables susceptibleHelicoverpa armigera to obtain resistance to Bacillus thuringiensis toxin Cry1Ac. BullEntomol Res 99:175–181.

33. Traut W (1997) A study of recombination, formation of chiasmata and synaptonemalcomplexes in female and male meiosis of Ephestia kuehniella (Lepidoptera). Genetica47:135–142.

34. Heckel DG, Gahan LJ, Liu YB, Tabashnik BE (1999) Genetic mapping of resistance toBacillus thuringiensis toxins in diamondback moth using biphasic linkage analysis.Proc Natl Acad Sci USA 96:8373–8377.

35. Elshire RJ, et al. (2011) A robust, simple genotyping-by-sequencing (GBS) approach forhigh diversity species. PLoS One 6:e19379.

36. Catchen JM, Amores A, Hohenlohe P, Cresko W, Postlethwait JH (2011) Stacks:Building and genotyping loci de novo from short-read sequences. G3 (Bethesda) 1:171–182.

37. Pearce SL, et al. (2017) Genomic innovations, transcriptional plasticity and gene lossunderlying the evolution and divergence of two highly polyphagous and invasiveHelicoverpa pest species. BMC Biol 15:63.

38. Li H, Durbin R (2009) Fast and accurate short read alignment with Burrows-Wheelertransform. Bioinformatics 25:1754–1760.

39. McKenna A, et al. (2010) The Genome Analysis Toolkit: A MapReduce framework foranalyzing next-generation DNA sequencing data. Genome Res 20:1297–1303.

40. Purcell S, et al. (2007) PLINK: A tool set for whole-genome association andpopulation-based linkage analyses. Am J Hum Genet 81:559–575.

41. Turner SD (May 14, 2014) qqman: An R package for visualizing GWAS results using Q-Q and Manhattan plots. bioRxiv:10.1101/005165.

42. van Dijk EL, Jaszczyszyn Y, Thermes C (2014) Library preparation methods for next-generation sequencing: Tone down the bias. Exp Cell Res 322:12–20.

43. Grabherr MG, et al. (2011) Trinity: Reconstructing a full-length transcriptome withouta genome from RNA-seq data. Nat Biotechnol 29:644–652.

44. Pertea G, et al. (2003) TIGR Gene Indices clustering tools (TGICL): A software systemfor fast clustering of large EST datasets. Bioinformatics 19:651–652.

Jin et al. PNAS | November 13, 2018 | vol. 115 | no. 46 | 11765

AGRICU

LTURA

LSC

IENCE

S

Dow

nloa

ded

by g

uest

on

Oct

ober

27,

202

0