The genome of Tetranychus urticae reveals herbivorous pest adaptations - Nature

Mites belong to the Chelicerata, the second largest group of terrestrial animals. Chelicerates represent a basal branch of arthropods. Subsequent to their origin in the Cambrian period, arthropods radiated into two lineages: the Chelicerata and the Mandibulata (comprising the Myriapoda and the Pancrustacea (which includes both crustaceans and insects))^1,2. Extant lineages of chelicerates include Pycnogonida, Xiphosura (horseshoe crabs) and Arachnida (a large group comprising scorpions, spiders and the Acari (ticks and mites))^3,4 (Supplementary Fig. 1.1). Within the Acari, T. urticae belongs to the Acariformes with the earliest fossils dating from the Lower Devonian period (410 million years ago). The Acari represent the most diverse chelicerate clade, with over 40,000 described species that exhibit tremendous variations in lifestyle, ranging from parasitic to predatory to plant-feeding. Some mites are of major concern to human health and include allergy-causing dust mites, scabies mites and mite vectors of scrub typhus⁵.

The two-spotted spider mite, Tetranychus urticae, is a cosmopolitan agricultural pest⁶ belonging to an assemblage of web-spinning mites. The name ‘spider’ highlights their ability to produce silk-like webbing used to establish a colonial micro-habitat, protect against abiotic agents, shelter from predators, communicate via pheromones and provide a vehicle for dispersion⁷.

Tetranychus urticae represents one of the most polyphagous arthropod herbivores, feeding on more than 1,100 plant species belonging to more than 140 different plant families including species known to produce toxic compounds. It is a major pest in greenhouse production and field crops, destroying annual and perennial crops such as tomatoes, peppers, cucumbers, strawberries, maize, soy, apples, grapes and citrus. The recent introduction of the related species Tetranychus evansi to Europe and Africa from South America demonstrates the invasive nature of these pests in global agriculture⁸. Computer modelling suggests that with intensifying global warming, the detrimental effects of spider mites in agriculture will markedly increase⁹ due to accelerated development at high temperatures.

Tetranychus urticae is known for its ability to develop rapid resistance to pesticides. Among arthropods it has the highest incidence of pesticide resistance¹⁰. Chemical control often causes a broad cross-resistance within and between pesticide classes, resulting in resistance to novel pesticides within 2–4 years. Many aspects of the biology of the spider mite, including rapid development, high fecundity and haplo-diploid sex determination, seem to facilitate rapid evolution of pesticide resistance. Control of multi-resistant mites has become increasingly difficult and the genetic basis of such resistance remains poorly understood¹¹.

As the first completed chelicerate genome, the comparison of the T. urticae genome with the genomes of insects and the crustacean Daphnia pulex expands the arthropod genetic toolkit. At the same time, the very compact T. urticae genome has unique attributes among arthropod genomes with remarkable instances of gene gains and losses. The completion of the T. urticae genome sequence opens new avenues for understanding the fundamentals of plant–herbivore interactions, developing novel pest-management strategies and producing new biomaterials on the nanometre scale.

The T. urticae genome (strain London) was sequenced (Sanger) to 8.05× coverage and assembled into 640 scaffolds covering 89.6 megabases (Mb) (Supplementary Notes 1, 2.1 and 2.2). 70,778 Sanger expressed sequence tag (EST) sequences from embryos, larvae, nymphs and adults were generated, and further complemented with RNA-seq data on matching samples (Supplementary Note 2.3). We identified 18,414 protein-coding gene models, of which 84% (15,397) are supported by EST (8,243), protein homology (11,433) and/or RNA-seq data (14,545) (Supplementary Note 2.4 and Supplementary Fig. 2.4.1). From alignments of ∼43-million paired-end Illumina reads from a second T. urticae strain (Montpellier) to the London sequence, 542,600 single nucleotide polymorphisms and small indels were predicted (Supplementary Note 2.5). The complete genome annotation of T. urticae is available at the BOGAS website¹². With an estimated genome size of about 90 Mb, the T. urticae genome is the smallest arthropod genome sequenced so far. The genomes of other chelicerates are much larger (565–7,100 Mb), with the unfinished genome of the tick Ixodes scapularis estimated at 2,100 Mb¹³. Multiple characteristics of the T. urticae genome correlate with its compact size: small transposable element content and microsatellite density, increased gene density and holocentric chromosomes (see Supplementary Note 3.1 for chromosomal features).

Transposable elements totalled 9.09 Mb (Supplementary Note 3.2), putting T. urticae together with D. pulex and Apis mellifera as arthropods with 10% or less of their genomes comprised of transposable elements. Long terminal repeat (LTR) retrotransposons, and in particular Gypsy-like elements, were the most abundant type of transposable elements. L1-like Long interspersed elements (LINEs), Tc1/Mariner-like DNA transposons, and Maverick (Polinton) elements were also detected (see Supplementary Table 3.2.1). Deep sequencing of small RNAs (∼19–30 nucleotides) across developmental stages (Supplementary Note 4.1) identified 226,829 unique RNAs that mapped to 676,266 different loci in the genome. The number of unique small RNA counts per size category shows a peak at 21 and 26 nucleotides. These two peaks include short interfering RNAs and Piwi-interacting RNAs, respectively, similar to what is observed in Drosophila melanogaster¹⁴. Their alignments to the genome indicate that both probably silence diverse transposable elements. Included among ∼21-nucleotide small RNAs are 52 predicted microRNAs (miRNAs). On the basis of the identity of their seed regions (nucleotides 2–7 of the miRNA sequence), the T. urticae miRNAs can be grouped into 43 families (Supplementary Note 4). Half of the predicted miRNAs were not conserved when compared to annotated miRNAs and available genomes of other arthropods¹⁵, suggesting that they might be T. urticae- or lineage-specific (Supplementary Tables 4.3.1–4.3.4).

The microsatellite density in the T. urticae genome is among the lowest observed for arthropods (Supplementary Note 3.3 and Supplementary Fig. 3.3.1), consistent with the expectation that repeat content of genomes typically scales with genome size. The T. urticae microsatellite classes have a distinct profile: mono-nucleotide repeats are virtually non-existent, and di-nucleotide repeats, normally the most abundant type of microsatellites, are found significantly less often than tri-nucleotides, as in Tribolium castaneum¹⁶. The gene density is twice as high compared to D. melanogaster, with 205 versus 92 genes per Mb, respectively. The mean number of exons per gene was low and similar to that found in D. melanogaster (∼3.8 exons per gene). The size distribution of introns was typically skewed with a mean intron size of 400 bp and a median of 96 bp (see Supplementary Note 3.4, Supplementary Fig. 2.4.3 and Supplementary Table 2.4.1). The holocentric nature of T. urticae chromosomes¹⁷ (the absence of centromeres and the diffuse nature of the kinetochores) is correlated with a lack of large tracts of gene-poor heterochromatin. The uniformly distributed gene density (Supplementary Note 3.1.1 and Supplementary Fig. 3.2.1) contrasts with the human body louse (Pediculus humanus, Phthiraptera, a hemimetabolous insect with a small genome), where 95% of the genes are concentrated in only 55 Mb of the 110-Mb genome¹⁸.

As the first completely sequenced and annotated chelicerate genome, the T. urticae genome expands the set of arthropod genomes beyond Pancrustacea and provides an important out-group for comparative genomics. Comparison of the coding gene repertoire of T. urticae with the arthropods T. castaneum, D. melanogaster, Nasonia vitripennis and D. pulex, the chordate Homo sapiens, and the cnidarian Nematostella vectensis (Fig. 1) resulted in 2,667 shared gene families (Supplementary Note 5.1). Almost 3,000 gene families are common to the arthropods sampled, whereas 5,038 gene families (8,329 genes) are unique to T. urticae (Supplementary Fig. 5.1.1). Of those, 622 gene families (1,398 genes) have homologues in species other than those listed above, most of which belong to other arthropods. Homologues of 74 gene families (93 genes) were found in the unfinished genomes of tick¹³ and/or Varroa destructor¹⁹ and are probably chelicerate, rather than specific to T. urticae. Therefore, 4,416 gene families (6,609 genes) were found to be unique to T. urticae. A gene gain/loss analysis (Fig. 1 and Supplementary Note 5.2) of these genomes showed a gain of about 700 new gene families in the lineage leading to T. urticae, plus almost 4,300 genes that are single copy (orphans). More than 1,000 gene families, still present in other arthropods, were lost in T. urticae. The 58 gene families that are significantly (z-score >2) expanded in T. urticae compared to the other arthropods are shown in Supplementary Note 5.2 and Supplementary Fig. 5.2.1.

At each time point (grey circles), the number of gains (+) and losses (−) of gene families is indicated as inferred by DOLLOP (black) and CAFÉ (red) programs. The inferred ancestral number of gene families, according to DOLLOP, is shown in green boxes.

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Tetranychus urticae is one of the most striking examples of polyphagy among herbivores and it has an unmatched ability to develop resistance to pesticides^6,10. We discovered that known gene families implicated in digestion, detoxification and transport of xenobiotics had a unique spider mite composition, and were often expanded when compared to insects (Supplementary Note 6.1). This included a threefold proliferation of cysteine peptidase genes, particularly C1A papain and the C13 legumain genes (Supplementary Table 6.1.11), consistent with proteolytic digestion based mostly on cysteine peptidase activity²⁰. Eighty-six cytochrome P450 (CYP) genes were detected in the T. urticae genome, a total number similar to insects but with an expansion of T. urticae-specific intronless genes of the CYP2 clan (Supplementary Table 6.1.2). The carboxyl/cholinesterases (CCEs) gene family contained 71 genes, with a single acetylcholinesterase gene (Ace1) but two new clades at the root of the neurodevelopmental class of CCEs, representing 34 and 22 CCEs, respectively (Supplementary Table 6.1.6). A notable case of expansion was found within the family of 32 glutathione S-transferases (GSTs) that include a group of 12 Mu-class GSTs that were, until now, believed to be vertebrate-specific (Supplementary Table 6.1.3). Finally, we discovered 39 multidrug resistance proteins belonging to the ATP-binding cassette (ABC) transporters (class C). The repertoire from this class of ABC transporters far exceeds the number (9–14) found in crustaceans, insects, vertebrates and nematodes (Supplementary Table 6.1.8). Few of the genes involved in detoxification had close insect homologues, and only four of the CYP genes could clearly be assigned as orthologues of insect and crustacean CYP genes.

The involvement of these gene families and their spider-mite-specific expansion in host plant adaptation is markedly illustrated by RNA-seq transcriptome profiling of spider mite feeding on its preferred host, bean (Phaseolus vulgaris), and on hosts to which the London strain is not adapted: Arabidopsis thaliana and tomato (Solanum lycopersicum) (Fig. 2) (Supplementary Notes 6.2). We found 24% of all genes to be differentially expressed upon host transfer (Fig. 2a–c); relative to bean, more genes were differentially expressed on tomato than on A. thaliana (Supplementary Note 6.2.4 and Supplementary Fig. 6.2.1), but responses were nonetheless correlated (Fig. 2b, c). Genes in the detoxification and peptidase families exhibited the most profound changes (Fig. 2a–c), with expression of nearly half of P450 genes affected by the host plant, including 19 of 39 genes in the intronless CYP392 family and the CYP389 family. These subfamilies are spider-mite-specific P450 expansions that define lineage-specific expansions²¹. This finding is unprecedented. In humans, only up to one-third of P450 genes are metabolizing xenobiotics²², and in D. melanogaster only one-third of the CYP genes are inducible by xenobiotics²³. The proportion of P450 genes responding to the chemical environment is much greater in the spider mite. Similar patterns were also found within other families (Fig. 2c). For GSTs and CCEs, the expression of Mu and Delta GSTs and the two spider-mite-specific CCE clades were most affected and about one-third of cysteine peptidases, the C1A papains and C13 legumains, were overexpressed after transfer to tomato. More than two-thirds of the CYP and GST genes affected by the host plant are present in clusters of (multiple) tandem duplicated genes. Co-regulation of the majority of tandem duplicates strongly indicates that the ancestral gene was already plant-responsive before duplication, and that a role in plant adaptation may have favoured duplicate retention.

a, A phylogeny of the cytochrome P450 (CYP) genes and heat map of the response of CYP genes to host transfer. Two-thirds of the genes that are tandemly duplicated or that form clusters (indicated by black vertical lines) are co-regulated. b, Global changes in gene expression after host shift. c, Fold changes of important gene family members in digestion and detoxification are colour coded. The analysis of differential expression (b and c) is with a 5% false discovery rate as assessed with RNA-seq data collected in biological triplicate (fold changes between mean values are plotted).

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Although these data indicate that spider-mite-specific expansion of known gene families contributes to the ability of spider mites to overcome host defences, many genes differentially regulated upon host transfer lack homology to genes of known function. Notably, among those with the most extreme expression fold-changes are genes that encode putative secreted proteins or lipid-binding proteins. Understanding extracellular binding and transport of small ligands is therefore likely to be important in further dissecting spider mite–plant interactions.

Our search for genes related to detoxification and digestion also revealed the existence and surprising expansion of intradiol ring cleavage dioxygenases, genes previously unreported from metazoan genomes but characteristic for bacteria and fungi²⁴. We annotated 16 functional genes in this family in T. urticae, whereas bacterial genomes usually carry only 1 to 7. They have an average sequence similarity of 43% with the homologue of Streptomyces avermitilis and share the conserved 2 His 2 Tyr non-haem iron(iii) binding site. These dioxygenases might have evolved to metabolize aromatic compounds found in plant allelochemicals. Other clear instances of lateral gene transfers include (1) the presence of a cobalamin-independent methionine synthase (MetE) gene with four predicted introns and up to 58% sequence identity to the MetE gene of soil Bacilli (this sequence has not previously been reported in any animal species); (2) two very similar levanase-encoding genes of probable bacterial origin that encode secreted exo-fructosidases upregulated upon feeding on tomato; and (3) a cyanate lyase-encoding gene that might be involved in feeding on cyanogenic plants (Supplementary Table 6.3.1).

We detected two clusters of carotenoid biosynthesis genes in T. urticae representing homologues of genes from zygomycete fungi and aphids. The latter are the only animal carotenoid biosynthesis genes known so far, thought to be derived from fungal genes by lateral gene transfer²⁵. The unique intron–exon structure of the spider mite and aphid genes and their clustering in phylogenetic analyses is strong evidence that the genes from fungi were transferred only once to arthropods (Fig. 3). The sequence and orientation of the two spider mite clusters indicate that they are the result of an ancient transfer followed by duplications, rearrangements and divergence. They also suggest that a second, more recent transfer occurred between a spider mite and an aphid ancestor, although the sequence of the two transfers remains speculative. Carotenoids are known to have a role in diapause induction in spider mites²⁶ and our findings indicate that they can also synthesize them.

The out-group comprises chimaeric assemblies (CSchim) of the closest bacterial sequences of cyclases and synthases. The T. urticae and Acyrthosiphon pisum sequences form a monophyletic group closely related to the zygomycete sequences. Evidence for a single lateral gene transfer event is also shown by the common intron positions in the cyclase/synthase (orange) and desaturase (green) genes (upper right panel). Two clusters of carotenoid biosynthesis genes are found in T. urticae: a tail-to-tail arrangement on scaffold 1 as seen in zygomycetes and aphids, and a more complex head-to-head (re)arrangement on scaffold 11 (bottom right).

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Hox genes are a conserved set of homeobox-containing transcription factors typically found clustered within the genome and used to establish region-specific identity during early development. The body plan of mites consists of an anterior prosoma and posterior opisthosoma and is further distinguished by an extremely reduced body plan presumably achieved through the fusion of segments (Supplementary Note 8 and Fig. 4b). The ancestral arthropod is predicted to have a Hox cluster with 10 genes²⁸. The T. urticae genome contains 8 of the canonical 10 genes. The ftz gene is present in duplicate, in two closely linked copies; orthologues of Hox3 and abdominal A (abdA) were not found (Fig. 4a). This is unusual among chelicerates: all 10 canonical Hox genes are present in the wandering spider²⁹. The absence of abdA in T. urticae correlates with the spider mite’s reduced opisthosomal segmentation. Consistent with the absence of abdA and a reduced opisthosoma, only two opisthosomal stripes of the segment polarity gene engrailed (typically expressed in each arthropod segment) are detected in the developing embryo (Fig. 4c), in contrast to five engrailed stripes detected in the opisthosoma of the wandering spider³⁰. Although numerous examples correlate morphological variation in arthropods with changes in Hox gene expression, this is the first example that correlates morphological evolution with the loss of a Hox gene within a fully sequenced Hox cluster.

a, T. urticae, T. castaneum and D. melanogaster Hox clusters. Gene sizes and intergenic distances are shown to scale. Dashed lines represent breaks in the cluster >1 Mb. In T. urticae, fushi tarazu and Antennapedia are present in duplicate whereas abdominal-A and Hox3/zerknullt are missing (red asterisk). b, Variable pressure scanning electron microscopy (SEM) image of adult T. urticae with two main body regions indicated: P, prosoma; O, opisthosoma. c, T. urticae engrailed (en) expression pattern. en transcripts are detected in five prosomal stripes that correspond to future pedipalpal (Pp), four walking leg (L1–L4) and two opisthosomal (O1 and O2) segments. Scale bars: b, 0.125 mm; c, 40 μm.

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Silk production in spider mites (Fig. 5a, b and Supplementary Note 9) represents a de novo evolution of silk-spinning relative to silk production in spiders⁷. Spiders typically spin silk from a complex glandular abdominal spinneret, whereas T. urticae uses paired silk glands connected to the mouth appendages (pedipalps)³¹. Seventeen fibroin genes were uncovered in the genome of T. urticae (Supplementary Table 9.1.1) encoding fibroins of unusually high (27–39%) serine content. We performed mechanical testing on fibres deposited by adult and larval mites with an atomic force microscope. This technique measures the Young’s modulus of the fibres, which is the ratio of applied stress (tension per cross-sectional area) to the resulting strain (fractional change in length) and describes the stiffness of the material. Young’s modulus was higher than or comparable to other natural materials (see Supplementary Table 9.1.2), but T. urticae silk fibres are thinner—54 ± 3 nm (adult silk, Fig. 5c) and 23.3 ± 0.9 nm (larval silk), that is, 435–185 times thinner—than the silk fibres of the spider Nephila clavipes³².

a, Spider mite colony on a bean plant forming characteristic silk webbing. b, SEM image of the spider mite larval silk filament (top), and atomic force microscopy (AFM) image of two larval spider mite silk filaments (bottom). c, Height profile of the adult spider mite silk filament obtained from the AFM image. Scale bars: a, 0.75 cm; b, 1 μm.

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Our analysis of the T. urticae genome also included nuclear receptors and neuropeptide genes, immunity-related genes and RNA interference, cuticle protein genes, and DNA methylation (Supplementary Notes 7.3 and 10–12).

The first complete genome of a chelicerate species provides the opportunity for a detailed phylogenomic analysis of arthropods, the most diverse group of animals on Earth. The T. urticae genome illustrates the specialized life history of this polyphagous herbivorous pest. Striking gene gains include lineage-specific expansions within detoxification gene families and lateral transfer of genes from fungi and bacteria that further expanded in T. urticae. The functional significance of these innovations is supported by the upregulation of many of these genes in response to feeding on less preferred host plants.

The genome of the two-spotted spider mite, together with the favourable biological features of the spider mite as a laboratory model including short generation time, easy rearing and tools for gene analysis and gene silencing³³, provide a novel resource for agriculture that should allow the dissection of pest–plant interactions and development of alternative tools for plant protection. Finally, evolutionary innovation in the process of T. urticae silk production expands the repertoire of potential chelicerate biomaterials (such as the well-known spider silk) with a natural biomaterial at the nanometre scale.

Primary accessions

GenBank/EMBL/DDBJ

Gene Expression Omnibus

Sequence Read Archive

Data deposits

Individual scaffolds of the T. urticae (London) genome are available through GenBank under accession numbers HE587301 to HE587940. The Illumina data for T. urticae strain Montpellier can be found in the Sequence Read Archive (SRA) database under accession numbers SRX030911 to SRX030913. RNA-seq data is available under Gene Expression Omnibus (GEO) super series number GSE32342.

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Author notes

1. Jeffrey A. Fawcett & Eric Bonnet

   Present address: Present addresses: Institut Curie, 26 rue d’Ulm, Paris 75248, France; INSERM, U900, Paris 75248, France ; Mines ParisTech, Fontainebleau 77300, France (E.B.); Graduate University for Advanced Studies, Hayama, Kanagawa 240-0193, Japan (J.A.F.).,

2. Miodrag Grbić, Thomas Van Leeuwen and Richard M. Clark: These authors contributed equally to this work.

Authors and Affiliations

1. Department of Biology, The University of Western Ontario, London N6A 5B7, Canada,

   Miodrag Grbić, Vojislava Grbić, Marc Cazaux, Marie Navarro, Vladimir Zhurov, Gustavo Acevedo & Anica Bjelica

2. Instituto de Ciencias de la Vid y el Vino (CSIC, UR, Gobiernode La Rioja), 26006 Logroño, Spain ,

   Miodrag Grbić & Vojislava Grbić

3. Department of Crop Protection, Faculty of Bioscience Engineering, Ghent University, B-9000 Ghent, Belgium,

   Thomas Van Leeuwen, Wannes Dermauw, Guy Smagghe, Masatoshi Iga, Olivier Christiaens & Luc Tirry

4. Department of Biology, University of Utah, Salt Lake City, 84112, Utah, USA

   Richard M. Clark & Edward J. Osborne

5. Department of Plant Systems Biology, VIB, Technologiepark 927, B-9052 Ghent, Belgium,

   Stephane Rombauts, Pierre Rouzé, Phuong Cao Thi Ngoc, Jeffrey A. Fawcett, Eric Bonnet, Cindy Martens, Guy Baele & Yves Van de Peer

6. Department of Plant Biotechnology and Bioinformatics, Ghent University, Technologiepark 927, B-9052 Ghent, Belgium,

   Stephane Rombauts, Pierre Rouzé, Phuong Cao Thi Ngoc, Jeffrey A. Fawcett, Eric Bonnet, Cindy Martens, Guy Baele & Yves Van de Peer

7. Department of Environmental Biology, Centro de Investigaciones Biológicas, CSIC, 28040 Madrid, Spain,

   Félix Ortego & Pedro Hernández-Crespo

8. Centro de Biotecnología y Genómica de Plantas,UPM-INIA, 28223 Madrid, Spain ,

   Isabel Diaz & Manuel Martinez

9. INRA, UMR CBGP (INRA/IRD/Cirad/Montpellier SupAgro), Campus international de Baillarguet, 34988 Montferrier-sur-Lez, France ,

   Maria Navajas

10. Instituto Gulbenkian de Ciência, 2781-901 Oeiras, Portugal ,

    Élio Sucena

11. Departamento de Biologia Animal, Universidade de Lisboa, Faculdade de Ciências, 1749-016 Lisbon, Portugal,

    Élio Sucena

12. Universidade de Lisboa, Faculdade de Ciências, Centro de Biologia Ambiental, 1749-016 Lisbon, Portugal ,

    Sara Magalhães

13. Department of Molecular and Cellular Biology, University of Arizona, Tucson, 85721, Arizona, USA

    Lisa Nagy & Ryan M. Pace

14. Department of Molecular Biology & Genetics, Johns Hopkins University School of Medicine, Baltimore, 21205, Maryland, USA

    Sergej Djuranović

15. Institut de Neurosciences Cognitives et Intégratives d’Aquitaine Université de Bordeaux 1, 33405 Talence, France ,

    Jan A. Veenstra

16. Centro Interdisciplinario de Neurociencia de Valparaíso, Facultad de Ciencias, Universidad de Valparaíso, Valparaíso 2360102, Chile ,

    John Ewer & Rodrigo Mancilla Villalobos

17. Department of Physics and Astronomy, The University of Western Ontario, N6A 5B7 London, Canada,

    Jeffrey L. Hutter & Stephen D. Hudson

18. Instituto de Catálisis y Petroleoquímica CSIC, Madrid, Spain; IMDEA Nanociencias, Facultad de Ciencias, Universidad Autonoma de Madrid, 28050 Madrid, Spain,

    Marisela Velez

19. IMDEA Nanociencias, Facultad de Ciencias, Universidad Autonoma de Madrid, 28050 Madrid, Spain,

    Marisela Velez

20. School of Biology, Georgia Institute of Technology, Atlanta, 30332, Georgia, USA

    Soojin V. Yi & Jia Zeng

21. Department of Biology, University of Texas at Arlington, Arlington, 76019, Texas, USA

    Andre Pires-daSilva

22. Universite de Toulouse, UPS, Centre de Biologie du Developpement, Universite Paul Sabatier, 31062 Toulouse, France; Centre National de la Recherche Scientifique, UMR 5547, Centre de Biologie du Developpement, 31062 Toulouse, France,

    Fernando Roch

23. Centre National de la Recherche Scientifique, UMR 5547, Centre de Biologie du Developpement, 31062 Toulouse, France,

    Fernando Roch

24. Westfälische Wilhelms University, Institute for Evolution and Biodiversity, Evolutionary Bioinformatics Group, Hüfferstrasse 1, D-48149 Münster, Germany ,

    Lothar Wissler

25. Department of Microbial and Molecular Systems, CMPG, K.U. Leuven, B-3001 Leuven, Belgium,

    Aminael Sanchez-Rodriguez

26. UPMC Univ Paris 06, UMR CNRS 7622, Equipe Biogenèse des signaux hormonaux, Case 29, 75005 Paris, France ,

    Catherine Blais

27. Department of Sustainable Organic Chemistry and Technology, Research Group EnVOC, Faculty of Bioscience Engineering, Ghent University, B-9000 Ghent, Belgium,

    Kristof Demeestere

28. Max Planck Institute for Developmental Biology, D-72076 Tübingen, Germany ,

    Stefan R. Henz

29. Department of Integrative Biology, University of Guelph, N1G 2W1 Guelph, Canada,

    T. Ryan Gregory

30. Boyce Thompson Institute for Plant Research, Ithaca, 14853, New York, USA

    Johannes Mathieu

31. Southern Crop Protection and Food Research Centre, Agriculture and Agri-Food Canada, N5V 4T3 London, Canada ,

    Lou Verdon

32. Fasteris SA, CH-1228 Plan-les-Ouates, Switzerland ,

    Laurent Farinelli

33. HudsonAlpha Institute for Biotechnology Huntsville, Alabama 35806, USA ,

    Jeremy Schmutz

34. DOE Joint Genome Institute, Walnut Creek, 94598, California, USA

    Jeremy Schmutz & Erika Lindquist

35. UMR 1301, INRA, CNRS and Université de Nice Sophia Antipolis, 06903 Sophia Antipolis, France ,

    René Feyereisen

Contributions

M.G. wrote the genome sequencing proposal. M.G. and V.G. generated DNA and RNA for sequencing. M.G., T.V.L., R.M.C., V.G., R.F. and Y.V.d.P. coordinated genome analysis and manuscript preparation. J.S. and E.L. coordinated genome assembly whereas P.R. and S.R. centralized and enabled the annotation process. All other authors are members of the spider mite genome sequencing consortium and contributed annotation, analyses or data to the genome project. M.G., T.V.L. and R.M.C. should be considered joint first authors. S.R., P.R. and V.G. should be considered joint second authors. M.G., R.F. and Y.V.d.P. should be considered joint corresponding authors.

Corresponding authors

Correspondence to Miodrag Grbić or Yves Van de Peer.

Competing interests

The authors declare no competing financial interests.

Supplementary Information

The file contains Supplementary Text, Supplementary Tables 1-12, Supplementary Figures 1-12 with legends and additional references (see Contents list for more details). (PDF 19025 kb)

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Grbić, M., Van Leeuwen, T., Clark, R. et al. The genome of Tetranychus urticae reveals herbivorous pest adaptations. Nature 479, 487–492 (2011). https://doi.org/10.1038/nature10640

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• Received: 30 June 2011

• Accepted: 17 October 2011

• Published: 23 November 2011

• Issue Date: 24 November 2011

• DOI: https://doi.org/10.1038/nature10640