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. 2025 Nov 17;7(1):100009. doi: 10.1016/j.abiote.2025.100009

Deciphering the dynamics of active autonomous terminal inverted repeat transposons in the plant kingdom

Ziye Huang a,b, Bicong Shi a, Li Huang a, Damon Lisch c, Xinyan Zhang a,⁎
PMCID: PMC12973397  PMID: 41940159

Abstract

Terminal inverted repeat (TIR) transposons are powerful drivers of genome evolution. However, a comprehensive understanding of their recent, lineage-specific activity across the plant kingdom has remained elusive. In this study, we developed a data-distillation pipeline to systematically identify recently active autonomous TIR transposons in 1007 plant genomes. Our analysis identified 3203 active clusters. The vast majority (93.3 ​%) of these clusters are maintained at low copy numbers, which suggests that robust host-mediated regulation restricts excessive proliferation. These TIR transposons exhibit striking heterogeneity and predominantly lineage-restricted diversification, with functional extinction being a common fate of TIR transposons across the plant kingdom. Among the identified active clusters, the Mutator-like element (MULE) superfamily was the most prevalent, accounting for 57.6 ​% of all TIR clusters. Furthermore, we uncovered extensive, previously uncharacterized intraspecific diversity through a genome analysis of four crop species, suggesting that single reference genomes substantially underestimate transposon dynamics. When we examined the molecular innovations that enable transposon success, we observed that MULE-encoded transposases possessed hypervariable termini that interact with accessory proteins. Using the hyperactive maize (Zea mays) MuDR element as a model, we obtained direct in vivo and in vitro evidence of an interaction between the transposase MURA and the accessory protein MURB, mapping the critical binding site to the N terminus of MURA. This atlas offers critical insights into transposon–host coevolution and provides a rich, species-specific toolkit for plant biotechnology.

Keywords: Accessory protein, Diversification, Lineage specificity, Plant genomes, TIR transposons

1. Introduction

Transposable elements (TEs) are dynamic DNA sequences that profoundly influence genetic diversity and genome evolution in cellular organisms [1]. These mobile genetic elements are broadly classified into two types based on their transposition mechanisms: Class I retrotransposons, which replicate through an RNA intermediate in a “copy-and-paste” procedure, and Class II DNA transposons, which excise and reinsert themselves via a “cut-and-paste” mechanism [2]. Among DNA transposons, terminal inverted repeat (TIR) transposons, which are structurally akin to prokaryotic insertion sequences, represent a particularly diverse group that has continuously shaped host genomes and provided genetic tools for advanced biotechnology [[3], [4], [5], [6]].

The potential for transposition activity is contingent upon the integrity of the transposon. Active autonomous TEs are self-sufficient genetic units that encode the transposase enzyme required for their own mobilization. Certain TEs also encode accessory proteins that assist with transposition. By contrast, non-autonomous elements can only transpose if an active, related transposon provides the necessary transposase in trans. Most active autonomous TEs act as genomic parasites, subject to strong purifying selection [7,8]. In some cases, active autonomous TEs have evolved to serve essential host functions, leading to a phenomenon known as host “addiction” where the host relies on the TE for critical aspects of its biology or for survival [9]. Thus, the ongoing mobility of autonomous TEs reflects a dynamic TE ecosystem within a genome and shapes the future trajectory of genomic evolution. This notion underscores the unique value of research into active autonomous TEs, in contrast to that of their non-autonomous variants.

TEs are ubiquitous in plant genomes, where they continuously shape genome diversity and can cause genomic instability. By generating variability in gene structure and expression, TEs provide the raw material for selective crop breeding, which ultimately enhances crop adaptability and influences agronomically important traits [[10], [11], [12]]. To counteract TE proliferation, plants have developed robust defense mechanisms, including transcriptional and post-transcriptional gene silencing mediated by small interfering RNAs [13,14]. This silencing mechanism intensifies with higher TE copy number, effectively limiting TE proliferation [15,16]. Despite host defenses, some active TEs persist by employing adaptive strategies. While canonical host–TE coevolution models emphasize transposon counter-defenses against epigenetic silencing, empirical support for anti-silencing mechanisms remains confined to context-specific cases. For instance, a CACTA transposon in rice (Oryza sativa) uses a microRNA to knock down the expression of DOMAINS REARRANGED METHYLTRANSFERASE 2 (OsDRM2), a gene essential for RNA-directed DNA methylation [17], while the VANDAL transposons in Arabidopsis (Arabidopsis thaliana) use rapidly evolving TIRs and accessory proteins to evade epigenetic silencing [18]. Broad-spectrum anti-silencing adaptations appear rare in plants, likely due to the risk of impairing reproductive success and hindering TE transmission through the germline [19]. Instead, some TEs maintain activity through targeted integration strategies. The maize (Zea mays) Mutator transposon, for instance, preferentially inserts near actively transcribed loci. This behavior is reminiscent of the P element in the fruit fly Drosophila melanogaster, suggesting that convergent evolution has led to similar adaptive mechanisms among eukaryotic TEs [20].

The mounting availability of high-quality plant genome assemblies over the past decade has revolutionized our understanding of transposon landscapes. Recent studies have offered valuable insights into mobilomes (i.e., the mobile genetic elements found within a genome) in plants on population-wide and species-spectrum scales [21,22]. However, detailed knowledge of active TIR-type TEs is largely confined to a small number of plant species and specific TE families [[23], [24], [25], [26]]. This restricted understanding of active TIR-type TEs has, in turn, hindered broader investigations into their ongoing contributions to genome variability and the complexity of TE–host interactions across the diverse plant kingdom.

Here, we present a curated collection of recently active autonomous TIR-type TEs from diverse plant lineages. Our study uncovered striking lineage-specific patterns and heterogeneity in the activity of recent TIR transposons, which shape the genomic landscape. We also investigated how the diversification and functionalization of Mutator-like element (MULE) TIRs have enhanced their adaptability. Together, our results provide a foundation for investigating TE–host interactions and developing biotechnological tools for diverse plants, including major crops.

2. Results

2.1. Recently active autonomous TIR transposons in plants exhibit lineage-specific dynamics

To explore the evolutionary dynamics of active autonomous TIR transposons across diverse plant lineages, we first annotated these genetic elements in the genomes of 1007 species belonging to the green lineage, ranging from algae to angiosperms, with each species represented by a single genome assembly (Fig. 1A; Table S1). Our approach defined “activity” as very recent divergence, leveraging stringent pairwise sequence dissimilarity thresholds and structural integrity (intact TIRs and target site duplications [TSDs]) as proxies for potential recent transposition. A critical challenge in such an analysis is that transposon sequence divergence is on a continuous spectrum, rendering attempts to categorically classify elements as “active” or “inactive” using discrete thresholds inherently arbitrary. Consistent with this conceptualization, we applied progressively higher maximum sequence dissimilarity thresholds of 0.1 ​%, 0.3 ​%, 1.0 ​%, and 3.0 ​%, which identified 561, 1170, 3203, and 6798 TIR TE clusters, respectively, spanning seven superfamilies (MULE, CACTA, hAT, PIF/Harbinger, Tc1/Mariner, Ginger, and Sola) (Fig. 1B). This result clearly highlights how threshold selection influences the sensitivity of transposon activity detection.

Fig. 1.

Fig. 1

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Pipeline tailored for the identification of recently proliferating autonomous TIR transposons across plant genomes. A ​Diagram illustrating the bioinformatic pipeline used to identify active autonomous TIR transposons. Colored boxes denote genes encoding transposases and accessory proteins within TIR transposons. ​B ​Number of active autonomous transposon clusters identified by pairwise sequence alignment under different dissimilarity thresholds. The 3.0 ​%, 1.0 ​%, 0.3 ​%, and 0.1 ​% dissimilarity thresholds correspond to 97.0 ​%, 99.0 ​%, 99.7 ​%, and 99.9 ​% sequence identity, respectively.

Given that active autonomous TIR transposons rely on the host DNA replication and repair machinery for their proliferation, we inferred that their sequence mutation rates are comparable to those of the overall plant genome (10−9 to 10−8 substitutions per site per year) [27]. Based on this inference, a 1 ​% divergence in sequence corresponds to an estimated divergence timeframe of 0.5–5.0 million years, which aligns with speciation events observed across numerous plant lineages [[28], [29], [30]]. Considering these insights, we selected a 1.0 ​% sequence dissimilarity threshold. This threshold yielded 3203 active clusters, which we classified as “recently active transposons” due to their intact architecture and minimal sequence divergence (Fig. 1, Fig. 2A; Table S2), while noting that direct evidence of ongoing transposition for most clusters will require experimental validation. Our pipeline accurately identified several known active TIR transposons in model organisms, including MuDR in maize and MCD7.9 (which has two copies with 99.1 ​% identity) in Arabidopsis [31,32]. Our pipeline effectively excluded more divergent elements, such as AtMu1 and AtMu2 of Arabidopsis, which have sequence identities of 98.8 ​% and 97.2 ​%, respectively, thus falling just below our stringent 99 ​% identity cutoff [31]. This observation demonstrates the high precision of our pipeline in selectively detecting the most recently amplified elements.

Fig. 2.

Fig. 2

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Genome-wide identification of active autonomous TIR transposons in 1007 plant species. A ​Summary table of active autonomous TIR transposons identified in the genomes of 1007 plant species. ​B ​Phylogenetic distribution of host species showing the type of TIR transposon and their abundance across 186 plant families. Darker shading indicates a higher value of the log2-transformed count of active TE clusters per species. The number of plant families in each clade is indicated in parentheses. ​C ​Distribution of copy number for major TIR transposon superfamilies. The copy number for each superfamily has been normalized such that the total number of clusters per superfamily is set to 1. ​D ​Violin plots showing the diversity of active TIR transposons as a function of genome size in their host species. Low variety, active autonomous TIR transposons from 1 to 2 superfamilies; high variety, active autonomous TIR transposons from 3 to 5 superfamilies. ​E ​Average number of active TE clusters per plant species across major angiosperm lineages.

Having defined clusters of recently active autonomous TIR transposons, we surveyed their distribution and population size to understand their survival dynamics. Nonvascular lineages (5.7 ​% of sampled species) accounted for only 0.9 ​% of the active transposon clusters detected (Fig. 2A). This underrepresentation suggests that these lineages more effectively control the accumulation of active TIR transposons. We observed massive expansion dynamics in vascular plants, which harbored more active autonomous TIR transposon clusters per species than algae and mosses (an average of 3.3 vs. 0.6 clusters) (Fig. 2A and B). Of note, 93.3 ​% (2988/3203) of active autonomous TIR transposon clusters maintained a low copy number (≤ 10 copies) of their autonomous elements, while high-copy clusters (> 10 copies) generally accounted for a minor proportion of three major superfamilies (CACTA: 12.4 ​%; hAT: 6.5 ​%; MULE: 4.3 ​%) (Fig. 2C). Only 0.9 ​% of all clusters (30/3203) were present as more than 50 copies (Table S2), reflecting strong selective pressure against excessive accumulation. Functional extinction, characterized by the absence of active transposon proliferation, was common across species: 42.7 ​% of species lacked recently proliferating autonomous TIR transposons from any of the seven superfamilies. Among the surveyed genomes, 17.3 ​% harbored active autonomous TIR transposons from 3 to 5 superfamilies (high variety), while 41.0 ​% had those from 1 to 2 superfamilies (low variety) (Fig. 2D). Species with greater TE diversity consistently had larger genomes (Fig. 2D). Of the high-variety cases, 49 species (4.8 ​%) hosted active elements from four superfamilies, while co-occurrence of five active TIR transposon types was restricted to three angiosperm species, wild sugarcane (Saccharum spontaneum), the Chinese medicine herb Kitagawia praeruptora, and barnyardgrass (Echinochloa crus-galli), suggesting exceptional genomic permissiveness in these plants. These results illuminate the lineage-specific dynamics underlying the expansion and persistence of autonomous TIR transposons in plant genomes.

Notably, active TE clusters from each superfamily exhibited distinct distribution patterns. Active autonomous MULE clusters accounted for 57.6 ​% (1846/3203) of overall active clusters, collectively dominating the active TIR transposon landscape across plant lineages, particularly in angiosperms (Table S2). Clusters of CACTA transposons, the second most abundant, represented 24.2 ​% (774/3203) of all clusters and displayed heightened activity in eudicots and monocots, but not in basal angiosperms (Fig. 2E). By contrast, Tc1/Mariner transposons, common in metazoans [4,33], showed limited mobilization across plant lineages (Fig. 2E). Ginger and Sola transposons were even rarer, comprising only 0.3 ​% (9/3203) of all clusters.

2.2. Extensive heterogeneity of active autonomous TIR transposons

To further evaluate the heterogeneity of active autonomous TIR transposons, we conducted all-versus-all sequence comparisons of consensus sequences from active autonomous TIR transposon clusters across each superfamily. Our analysis revealed predominantly low pairwise sequence identity across all TIR superfamilies (Fig. 3A), indicating that most recently active transposons have evolved independently within their host lineages. We also identified 76 interspecific near-identical pairs of active clusters (≥ 99.0 ​% sequence identity) (Fig. 3B; Table S3), 66 of which were present in closely related species. We detected 20 of these pairs of clusters within species sharing homoeologous subgenomes (one or both being polyploid genomes), while 33 were present in the genomes of domesticated crop species and their wild relatives (Table S3), suggesting that autonomous TIR transposons have been maintained during polyploid speciation and crop domestication. Among the remaining ten pairs, nine were in related plant species within the same genus, and the last pair spanned different genera within a plant family (Virginia strawberry [Fragaria virginiana] and silverweed [Potentilla anserina]) (Fig. 3B; Table S3). BLAST searches confirmed the widespread distribution of this TE cluster, identifying highly similar (> 97 ​% identity), full-length sequences in four species across the genera Fragaria, Potentilla, and Dasiphora (Fig. 3C), which is consistent with possible vertical inheritance from a common ancestor.

Fig. 3.

Fig. 3

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Interspecies sequence heterogeneity of active autonomous TIR transposons. A ​Heatmap visualization of the percentage identity in an all-vs-all comparison of the orientation-corrected consensus sequences from active TIR transposon clusters. ​B ​Pie chart describing 76 paired plant species harboring highly similar active autonomous TIR clusters. ​C ​Phylogeny of a clade of highly similar ​PIF/Harbinger ​elements across genera in the ​Rosaceae. Bootstrap support values ​≥ ​70 are indicated.

To assess the evolutionary constraints imposed on this element, we compared its sequence divergence with the average synonymous substitution rate (Ks) of 12 orthologous genes between F. virginiana and P. anserina. The average Kimura distance of the TE cluster was exceptionally low (0.0049), nearly 50-fold lower than the average gene Ks of 0.2397 for the set of 12 genes analyzed here. This lower degree of divergence suggests that these TE clusters may be subject to strong purifying selection; however, the possibility of frequent horizontal transfer cannot be ruled out based on current evidence.

To evaluate whether lineage-specific profiles reflect species-wide diversity, we analyzed four geographically distinct accessions for each of four representative species (maize, rice, potato [Solanum tuberosum], and field mustard [Brassica rapa]) (Table S4). Through similarity-based clustering of TIRs, we resolved four canonical active TIR families (MULE, CACTA, hAT, and PIF/Harbinger) into 102 discrete clusters (Fig. 4; Table S5). Phylogenetic conservation was strictly species-specific: we detected no clusters with interspecific conservation across these four sets of crop genomes surveyed. An analysis within each species revealed remarkable diversification, with 63.7 ​% (65/102) of all clusters exhibiting strict restriction by genotype and only 2.9 ​% (3/102) displaying conservation across all four accessions within a species (Fig. 4). This observation brings to light a wealth of active TIR transposons in crops, and also advocates for multi-accession surveys to adequately capture intraspecific TE heterogeneity and dynamics.

Fig. 4.

Fig. 4

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Evaluation of intraspecies diversification of active TIR transposon clusters across accessions of four plant species. Each column represents a unique TIR TE cluster. Circle area reflects population size within the genome of one host accession.

2.3. Structural diversity and protein modularity in MULE transposons of plants

We investigated the sequence diversity of the proteins encoded by the active TIR clusters. Both structural conservation and structural diversification were present in the transposases encoded by each superfamily, as shown in multiple amino acid sequence alignments of transposases predicted from the consensus sequences of active clusters (Fig. 5A and Fig. S1A). Of the non-transposase proteins encoded by active transposons, 52.3 ​% contained identifiable conserved domains. In addition to recurrent domains (e.g., Zf-GRF, Peptidase_C48, and NAM-associated), most domains were present in few proteins encoded by active transposons, with 335 distinct PFAM domains appearing as singletons (Fig. S1B; Table S6).

Fig. 5.

Fig. 5

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The N-terminal peptide of the transposase encoded by MuDR interacts with the accessory protein. A ​Structural alignment of transposases encoded by active ​MULE ​transposons, highlighting conserved domains. Each transposase is represented by a horizontal line, with lines vertically aligned according to the locations of the conserved transposase domains. TEs were classified based on the combinations of identifiable conserved domains. ​B ​Length of non-conserved N-terminal regions in transposases encoded by the different clades of ​MULE ​elements. ​C ​Diagram of the active ​MuDR ​transposon. ​D ​Co-immunoprecipitation (Co-IP) assay showing that the first 209 ​amino acids of MURA interact with MURB. The indicated constructs were co-infiltrated into the leaves of ​Nicotiana benthamiana ​plants and total proteins were extracted prior to Co-IP with an anti-FLAG antibody. ​E ​GST pull-down assay confirming the binding of MURB to the MURA N terminus. GST or GST-MURA was immobilized onto glutathione beads before incubation with purified recombinant MBP or MBP-MURB. Bead-bound proteins were probed by immunoblotting with ​anti-GST and ​anti-MBP antibodies. ​F ​Split-luciferase assay verifying that the N terminus of MURA interacts with MURB. A ​N. benthamiana ​leaf was co-infiltrated with the indicated constructs, sprayed with D-luciferin, and imaged for luciferase activity, shown here on a false color scale.

To elucidate the adaptations of MULE transposons to the environment of plant genomes, we conducted a detailed analysis of the structure and characteristics of proteins encoded by active MULE transposons. Structurally, recently active MULE transposons had the longest TIRs among the families studied, with an average length of 200 bp and considerable variation (range: 20–1479 bp), in contrast to other families with shorter TIRs (mean: 28–34 bp) (Fig. S2A). Additionally, the target site sequences of active MULE elements were predominantly random, showing only a marginal preference for AT-rich sequences (Fig. S2B). This observation contrasted with that of certain families, such as the Mutator elements in the maize and rice genomes, which favor either GC-rich or AT-rich sequences [31,34,35]. These observations suggest that while the MULE superfamily exhibits broad targeting flexibility, specific families have likely evolved more refined targeting mechanisms. Beyond their structural features, MULE-encoded transposases exhibited distinct characteristics. Most MULE transposases had extended N- and C-terminal extensions not present in those of other families (Fig. 5A and Figs. S1A, C, and D). Notably, MURA-type transposases within MULE elements featured substantially longer N-terminal extensions than JITA-type variants (Fig. 5B). Furthermore, a subset of MURA-type transposases across multiple independent lineages contained a Plant Mobile Domain (PMD) in their C-terminal region, indicating a potential role for the ends of transposases in the adaptation of MULE transposons to plant genomes.

To explore the potential protein modules encoded by MULEs, we examined the proteins encoded by the MuDR transposon in maize, a well-established model for MULEs due to its hyperactive transposition [36]. MuDR encodes the MURA transposase and the accessory protein MURB (Fig. 5C), both critical for heritable de novo insertions [37], although direct biochemical evidence of their interaction has been lacking. Functional characterization of MURA has been hindered due to the cytotoxicity of the encoding gene during cloning in bacteria [38]. The toxicity persisted despite the codon optimization in our analysis. Through random mutagenesis, we identified an R724K substitution that successfully lowered the cytotoxicity of the protein to a level suitable for cloning, as demonstrated by the successful propagation of plasmids carrying the modified gene in the bacterial hosts. Leveraging this mutant, we performed a co-immunoprecipitation assay, using total protein extracts from the leaves of Nicotiana benthamiana plants that had been infiltrated with constructs encoding MURAR724K-GFP (a fusion of MURAR724K and green fluorescent protein) and purified MBP-MURB (a fusion of maltose-binding protein and MURB) expressed in E. coli. Following immunoprecipitation with an anti-GFP antibody, we detected MURB among the co-precipitated proteins, indicative of an-interaction between MURA and MURB (Fig. S3). A truncation analysis mapped the interaction interface to the N-terminal region of MURA (residues 1–209) (Fig. 5D), a result corroborated by split-luciferase and glutathione S-transferase (GST) pull-down assays (Fig. 5E and F). These results demonstrate the critical role of the variable N-terminal domain of the transposase MURA in mediating interactions with the accessory protein MURB, which is encoded by active MULEs.

3. Discussion

In contrast to prior studies that either examined TEs from a population within a given species or broadly surveyed TEs across plant lineages [21,22], our study provides previously missed insights into the dynamics of recently active autonomous TIR transposons. Our specialized data-distillation pipeline allowed us to generate a curated catalog of the recently proliferating autonomous TIR transposons from 1007 high-quality genome assemblies across the green lineage, from algae to angiosperms. Our data reveal lineage-specific patterns, heterogeneous distribution, and unique structural adaptations, emphasizing rapid TIR transposon diversification and adaptability. The inherent versatility of these endogenous active TIR transposons, previously demonstrated in both plant and animal systems, positions them as highly promising tools for mutagenesis and gene delivery [3,4]. Furthermore, the host specificity of TIR transposons identified in our dataset provides a valuable resource for developing tailored toolkits for targeted plant species, minimizing the need for extensive optimization.

Our observations reveal the diverse activity of lineage-specific autonomous TIR transposons across genomes in the green lineage and provide insights into TE activity and host permissiveness, factors that ultimately dictate the long-term expansion and extinction of TEs [39]. We uncovered a patchy landscape of recent TIR transposon activity throughout the green lineage, indicating that functional extinction is a common fate for these elements. Interestingly, some plant lineages simultaneously harbor recently proliferating TIR elements from three or more superfamilies, suggesting that these lineages have a permissive genomic environment that allows TIR transposons to thrive. We determined that the vast majority of recently active autonomous TIR clusters are maintained at low copy numbers. This observation supports an evolutionary model in which restrained proliferation minimizes fitness costs to the host, thereby allowing for elevated adaptation [40]. While maintaining a low copy number is a prevalent strategy, the distribution of active TIR transposons is far from uniform. The striking rarity of active clusters with high copy-number highlights the strong purifying selection against uncontrolled TIR TE proliferation in most plant genomes. This rarity also suggests that underlying mechanisms enable temporary escapes from host regulatory control.

Our work revealed significant heterogeneity in active TIR transposons both across and within plant species. Pairwise comparisons of active transposon clusters in our dataset suggest that vertical transmission is their predominant mode of inheritance. Furthermore, our survey of four representative crop species highlights substantial TE heterogeneity within each species, underscoring the need for multi-genome surveys to enhance our understanding of the abundance and diversity of active transposons. These observations provide valuable insights for leveraging a broader range of accession-specific active TIR transposons in crop genomes to advance future breeding and genetic engineering efforts.

Among the seven TIR superfamilies, MULE transposons have emerged as the most successful in colonizing recent plant genomes, as revealed by this survey. Recently active MULE elements possess exceptionally long TIRs, with some extending up to 1.5 ​kb. These elongated TIRs likely facilitate the formation of chimeric elements and promote genetic exchange among MULE elements [41]. Additionally, we observed hypervariable termini in MULE-encoded transposases, accompanied by a diverse array of associated accessory proteins. This variability suggests lineage-specific functional innovations and adaptations, some of which are conserved across distinct lineages. These observations align with prior systematic evaluations of Mutator element evolution across diverse eukaryotic lineages and the divergence in the evolutionary speed of MuDR-like Abermu elements across different open reading frames [42,43]. For instance, the PMD domain, present in multiple active clusters, is likely selected for its role in counteracting H3K27me3-mediated gene silencing through chromatin remodeling [[44], [45], [46]].

Our observations provide empirical evidence for modular interactions between the ends of transposases and accessory proteins, as exemplified by the hyperactive maize MuDR element. Although earlier yeast two-hybrid assays failed to detect an interaction between MURA and MURB [47], we were able to validate an interaction in vivo and in vitro between the N-terminal region of the transposase MURA and the accessory protein MURB. These observations align with genetic evidence showing that the loss of MURB function markedly inhibits the transposition of Mutator elements, likely by disrupting the integration step [37]. Given the floral meristem–specific expression pattern of MURB [37,48,49], this interaction suggests a specialized mechanism for fine-tuning transposition activity in response to host developmental cues through a tethering mechanism. Although direct interactions between transposases and accessory proteins have been documented in the PIF/Harbinger and CACTA superfamilies [50,51], such interactions have been newly identified in the MULE superfamily, revealing a convergent evolutionary pattern among TIR superfamilies. Collectively, these observations highlight the pivotal roles of structural variation and modular functionality in driving the lineage-specific adaptation of MULE transposons.

Taken together, our study delivers a meticulously curated dataset of TEs, revealing a dynamic landscape of active autonomous TIR transposons across the genomes of the green lineage and providing evidence for genome-specific adaptations to their transposition activity. Building on this work, future research should dissect the dynamics of active TIR TEs at the subgenomic level, an area not addressed in detail here. Additionally, future work should address the biases of TE dynamics due to limited genome availability by applying expanded phylogenetic sampling, particularly in basal plant lineages. Although our work focused on TIR transposons in plants, the principle of diversification-driven persistence uncovered here may extend to a wide range of TEs across diverse eukaryotic lineages.

4. Materials and methods

4.1. Genome dataset and transposon annotation

Chromosome-level and complete assemblies from the genomes of 1007 plant species (contig N50 value ​> ​12 ​kb) were retrieved from the National Center for Biotechnology Information (NCBI, US) and the National Genomics Data Center (NGDC, China). TIR transposons were annotated using MITE-Tracker (TSD length: 2–15 bp; element length: 2000–12000 bp) [52]. Transposase-encoding autonomous elements were identified through six-frame translation of the retrieved sequences, followed by screening for the relevant conserved domains using hmmsearch (Pfam-A database, version 37.0; e-value < 1e−5) [53], retaining elements encoding proteins with conserved transposase domains from 17 transposon superfamilies (MULE, CACTA, hAT, PIF/Harbinger, Tc1/Mariner, Ginger, Sola, Merlin, Kolobok, Zator, Zisupton, Transib, PiggyBac, P, Academ, Novosib, and Dada). For Ginger, Sola, Academ, Novosib, and Dada, domain profiles were generated using HMMER software based on multiple sequence alignments of the conserved domain regions from their respective transposases. Candidate TIR pairs with high full-length sequence identity (≥ 99.9 ​%, ≥ 99.7 ​%, ≥ 99.0 ​%, and ≥ 97.0 ​%) and < 80 ​% flanking sequence identity (using pairwise global alignment) were subjected to a sensitivity test. These candidates were clustered through a disjoint-set algorithm (≥ 90 ​% identity threshold) with iterative pairwise alignment merging [54]. Specifically, any two TE pairs sharing a common TE were merged into a larger set, with additional pairs incorporated if specific TEs connected them to the existing group. ‘Putatively recently active’ autonomous TIR transposon clusters were operationally defined as those maintaining ​≥ ​two structurally intact copies in a given genome with a sequence identity ≥ 99.0 ​%, indicative of recent proliferation and minimal sequence decay.

For long TIRs, sequence similarity was analyzed exclusively in their terminal 70-bp regions. HMM-based gene structure prediction and the protein sequences encoded by autonomous transposons were analyzed using Fgenesh with Arabidopsis-optimized parameters [55]. Plant family phylogeny was reconstructed from LifeMap subtree topology (https://lifemap.univ-lyon1.fr/) [56]. Comparative analysis employed Hiplot Pro for violin and boxplot visualizations (https://hiplot.com.cn/). Orientation-corrected consensus sequences from active TIR transposon clusters were subjected to all-vs-all comparisons where global pairwise alignments were performed using MAFFT v7.520 with accelerated parameters (--retree 1 --maxiterate 0) [57]. Source data for aligned transposase sequences, used to create domain profiles with hmmbuild, along with custom scripts for transposable element annotation, have been deposited in GitHub (https://github.com/zhang-caas/TIR_cluster).

4.2. Clustering and pairwise comparison of TIR transposons

To analyze intra-species TIR transposon diversity, a two-step sequence similarity–based clustering approach was employed. First, TIR sequences were compared pairwise by aligning the first 70 bp of the left end of the TIRs and the last 70 bp of the right end of the TIRs (via pairwise global alignment). Pairs with ≥ 90 ​% identity over both TIRs, or in the reverse-complement orientation, were marked. These pairwise relationships were then aggregated into distinct clusters using a disjoint-set data structure. Custom scripts for TE clustering analysis have been deposited in GitHub (https://github.com/zhang-caas/TIR_cluster).

4.3. Calculation of evolutionary rates of genes and TIR transposons

To establish a benchmark for the rate of neutral evolution, the coding sequence of 12 orthologous genes from Fragaria vesca and Potentilla anserina was analyzed. The synonymous substitution rate (Ks) was calculated for each pair using the Nei-Gojobori method [58], and the arithmetic mean of these Ks values was computed for use in subsequent comparisons. To quantify the evolutionary divergence of TIR elements, the average pairwise genetic distance was calculated using the Kimura 2-Parameter model [59]. For each pair of TIR elements, the Kimura distance was determined based on aligned sequences. The final evolutionary rate was derived by calculating the arithmetic mean of all valid pairwise distances.

4.4. Molecular cloning

All genes cloned in this study were derived from cDNA prepared from total RNA extracted from a Mutator-active maize (Zea mays) line described in a previous study [37]. The MURAR724K mutant (full-length, aa 1–823) harbors an R724K substitution. For co-immunoprecipitation (Co-IP) assays, the coding sequences of MURAR724K and MURA1-209aa were cloned in-frame and upstream of the sequence encoding eGFP in the pEAQ vector, while the coding sequence of MURB was placed in-frame and upstream of the sequence encoding a C-terminal 3 ​× ​FLAG tag in the pEAQ vector. In pull-down assays, the coding sequence of MURA1-209aa was cloned in-frame and downstream of the sequence encoding GST in the pGEX-6P vector, while the coding sequence of MURB was placed in-frame and upstream of the sequence encoding MBP in the pMAL-p2x vector. For split-luciferase assays, the coding sequence of MURA1-209aa was cloned in-frame and upstream of the sequence encoding nLUC, while the coding sequence of MURB was placed in-frame and downstream of the sequence encoding cLUC, in the pCAMBIA 1300-nLUC and pCAMBIA 1300-cLUC vectors, respectively. All constructs were assembled by homologous recombination using the ClonExpress II One Step Cloning Kit (Vazyme).

4.5. Co-immunoprecipitation (Co-IP)

Leaf tissue from Nicotiana benthamiana plants (250 ​mg) expressing the appropriate constructs encoding GFP-tagged and FLAG-tagged proteins was homogenized in 750 ​μL Co-IP buffer (50 ​mM Tris-HCl pH 7.5, 150 ​mM NaCl, 1 ​% [v/v] NP-40, 1 ​mM DTT) containing 1 ​× ​cOmplete protease inhibitor cocktail (Roche). The supernatants were incubated with GFP-Trap Agarose beads for 2 ​h at 4 ​°C. The beads were then washed with Co-IP buffer, incubated with recombinant purified MBP-tagged proteins for 2 ​h, and washed again. For co-expression assays with protein extracts prepared from leaf tissue expressing constructs encoding GFP-tagged and FLAG-tagged proteins, the beads were washed after incubation with the protein extracts. Bead-bound proteins were eluted in SDS-loading buffer and analyzed by immunoblotting with anti-GFP or anti-FLAG antibodies (Abclonal, 1:5000 dilutions).

4.6. GST pull-down assay

GST-tagged and MBP-tagged proteins were produced in Escherichia coli BL21. The cell pellet was collected and lysed in Tris-HCl buffer (pH 7.5, 150 ​mM NaCl, 1 ​mM DTT) containing 1 ​× ​protease inhibitor cocktail, and the mixture clarified by centrifugation (6000×g, 6 ​min, 4 ​°C). GST-tagged proteins were immobilized on GSTSep Glutathione Agarose Resin (Yeasen) and incubated with purified MBP-tagged proteins (2 ​h, 4 ​°C). After extensive washing with Tris-based wash buffers, bound complexes were eluted with SDS buffer and detected by immunoblotting using anti-GST or anti-MBP antibodies (Abclonal, 1:5000 dilutions).

4.7. Split-luciferase assay

Cell cultures from pairs of Agrobacterium tumefaciens (strain GV3101) positive colonies carrying each of two split-luciferase plasmids were resuspended in infiltration buffer (10 ​mM MgCl2, 10 ​mM MES pH 5.6, 150 ​μM acetosyringone), mixed in a 1:1 (v/v) ratio to an optical density (OD600) of 0.8–1.0, and co-infiltrated into the leaves of N. benthamiana plants. After 36 ​h, D-luciferin substrate (Macklin, catalog number: D807792) was sprayed onto the abaxial side of the leaves; following dark adaptation for 1 ​min, leaves were imaged using a chemiluminescence detection system (Lumazone PyLoN2048B, Roper Scientific).

CRediT authorship contribution statement

Ziye Huang: Writing – original draft, Methodology, Investigation, Formal analysis. Bicong Shi: Validation, Methodology, Investigation. Li Huang: Methodology, Investigation, Funding acquisition. Damon Lisch: Writing – review & editing, Validation, Formal analysis. Xinyan Zhang: Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Resources, Project administration, Investigation, Funding acquisition, Formal analysis, Data curation, Conceptualization.

Declaration of competing interest

On behalf of all authors, the corresponding author states that there is no conflict of interest.

Acknowledgements

The authors express their gratitude to Dr. Meixia Zhao and Dr. Yu Zhang for their insightful discussions and generous assistance with the manuscript. This work was supported by National Key Research and Development Program of China (2023YFF1001100 and 2023YFF1000400), National Natural Science Foundation of China (32070552 and 32400254), and Guangdong Major Project of Basic and Applied Basic Research (2021B0301030004).

Footnotes

Appendix A

Supplementary data to this article can be found online at https://doi.org/10.1016/j.abiote.2025.100009.

Appendix A. Supplementary data

The following is the supplementary data to this article:

Multimedia component 1
mmc1.xlsx (7.1MB, xlsx)
Multimedia component 2
mmc2.pdf (262KB, pdf)

Data availability

Information regarding genome assemblies and active autonomous TE clusters is available in the online Supplementary Data.

References

  • 1.Wells J.N., Feschotte C. A field guide to eukaryotic transposable elements. Annu Rev Genet. 2020;54:539–561. doi: 10.1146/annurev-genet-040620-022145. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Wicker T., Sabot F., Hua-Van A., Bennetzen J.L., Capy P., Chalhoub B., et al. A unified classification system for eukaryotic transposable elements. Nat Rev Genet. 2007;8(12):973–982. doi: 10.1038/nrg2165. [DOI] [PubMed] [Google Scholar]
  • 3.Liu P., Panda K., Edwards S.A., Swanson R., Yi H., Pandesha P., et al. Transposase-assisted target-site integration for efficient plant genome engineering. Nature. 2024;631(8021):593–600. doi: 10.1038/s41586-024-07613-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Zhang T., Tan S., Tang N., Li Y., Zhang C., Sun J., et al. Heterologous survey of 130 DNA transposons in human cells highlights their functional divergence and expands the genome engineering toolbox. Cell. 2024;187(14):3741–3760 e30. doi: 10.1016/j.cell.2024.05.007. [DOI] [PubMed] [Google Scholar]
  • 5.Feschotte C., Pritham E.J. DNA transposons and the evolution of eukaryotic genomes. Annu Rev Genet. 2007;41:331–368. doi: 10.1146/annurev.genet.40.110405.090448. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Li Q., Wang Y., Hou Z., Zong H., Wang X., Zhang Y.E., et al. Genome editing in plants using the TnpB transposase system. aBIOTECH. 2024;5(2):225–230. doi: 10.1007/s42994-024-00172-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Horvath R., Minadakis N., Bourgeois Y., Roulin A.C. The evolution of transposable elements in Brachypodium distachyon is governed by purifying selection, while neutral and adaptive processes play a minor role. eLife. 2024;12 [Google Scholar]
  • 8.Langmüller A.M., Nolte V., Dolezal M., Schlötterer C. The genomic distribution of transposable elements is driven by spatially variable purifying selection. Nucleic Acids Res. 2023;51(17):9203–9213. doi: 10.1093/nar/gkad635. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Chang N.C., Wells J.N., Wang A.Y., Schofield P., Huang Y.C., Truong V.H., et al. Gag proteins encoded by endogenous retroviruses are required for zebrafish development. Proc Natl Acad Sci U S A. 2025;122(18) [Google Scholar]
  • 10.Yang Q., Li Z., Li W., Ku L., Wang C., Ye J., et al. CACTA-like transposable element in ZmCCT attenuated photoperiod sensitivity and accelerated the postdomestication spread of maize. Proc Natl Acad Sci U S A. 2013;110(42):16969–16974. doi: 10.1073/pnas.1310949110. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Bhattacharyya M.K., Smith A.M., Ellis T.H., Hedley C., Martin C. The wrinkled-seed character of pea described by mendel is caused by a transposon-like insertion in a gene encoding starch-branching enzyme. Cell. 1990;60(1):115–122. doi: 10.1016/0092-8674(90)90721-p. [DOI] [PubMed] [Google Scholar]
  • 12.Castanera R., Morales-Diaz N., Gupta S., Purugganan M., Casacuberta J.M. Transposons are important contributors to gene expression variability under selection in rice populations. eLife. 2023;12:RP86324. doi: 10.7554/eLife.86324. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Lisch D. Epigenetic regulation of transposable elements in plants. Annu Rev Plant Biol. 2009;60:43–66. doi: 10.1146/annurev.arplant.59.032607.092744. [DOI] [PubMed] [Google Scholar]
  • 14.Creasey K.M., Zhai J., Borges F., Van Ex F., Regulski M., Meyers B.C., et al. miRNAs trigger widespread epigenetically activated siRNAs from transposons in Arabidopsis. Nature. 2014;508(7496):411–415. doi: 10.1038/nature13069. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Borges F., Martienssen R.A. The expanding world of small RNAs in plants. Nat Rev Mol Cell Biol. 2015;16(12):727–741. doi: 10.1038/nrm4085. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Mari-Ordonez A., Marchais A., Etcheverry M., Martin A., Colot V., Voinnet O. Reconstructing de novo silencing of an active plant retrotransposon. Nat Genet. 2013;45(9):1029–1039. doi: 10.1038/ng.2703. [DOI] [PubMed] [Google Scholar]
  • 17.Nosaka M., Itoh J., Nagato Y., Ono A., Ishiwata A., Sato Y. Role of transposon-derived small RNAs in the interplay between genomes and parasitic DNA in rice. PLoS Genet. 2012;8(9) [Google Scholar]
  • 18.Sasaki T., Ro K., Caillieux E., Manabe R., Bohl-Viallefond G., Baduel P., et al. Fast co-evolution of anti-silencing systems shapes the invasiveness of Mu-like DNA transposons in eudicots. EMBO J. 2022;41(8) [Google Scholar]
  • 19.Haig D. Transposable elements: self-Seekers of the germline, team-players of the soma. Bioessays. 2016;38(11):1158–1166. doi: 10.1002/bies.201600125. [DOI] [PubMed] [Google Scholar]
  • 20.Zhang X., Zhao M., McCarty D.R., Lisch D. Transposable elements employ distinct integration strategies with respect to transcriptional landscapes in eukaryotic genomes. Nucleic Acids Res. 2020;48(12):6685–6698. doi: 10.1093/nar/gkaa370. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Huang Y., Sahu S.K., Liu X. Deciphering recent transposition patterns in plants through comparison of 811 genome assemblies. Plant Biotechnol J. 2025;23(4):1121–1132. doi: 10.1111/pbi.14570. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Li X., Dai X., He H., Lv Y., Yang L., He W., et al. A pan-TE map highlights transposable elements underlying domestication and agronomic traits in Asian rice. Natl Sci Rev. 2024;11(6):nwae188. doi: 10.1093/nsr/nwae188. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Hirochika H., Sugimoto K., Otsuki Y., Tsugawa H., Kanda M. Retrotransposons of rice involved in mutations induced by tissue culture. Proc Natl Acad Sci U S A. 1996;93(15):7783–7788. doi: 10.1073/pnas.93.15.7783. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Lisch D., Chomet P., Freeling M. Genetic characterization of the mutator system in maize: behavior and regulation of Mu transposons in a minimal line. Genetics. 1995;139(4):1777–1796. doi: 10.1093/genetics/139.4.1777. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Fedoroff N., Wessler S., Shure M. Isolation of the transposable maize controlling elements Ac and Ds. Cell. 1983;35(1):235–242. doi: 10.1016/0092-8674(83)90226-x. [DOI] [PubMed] [Google Scholar]
  • 26.Chen J., Lu L., Benjamin J., Diaz S., Hancock C.N., Stajich J.E., et al. Tracking the origin of two genetic components associated with transposable element bursts in domesticated rice. Nat Commun. 2019;10(1):641. doi: 10.1038/s41467-019-08451-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Wang Y., Obbard D.J. Experimental estimates of germline mutation rate in eukaryotes: a phylogenetic meta-analysis. Evol Lett. 2023;7(4):216–226. doi: 10.1093/evlett/qrad027. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Good-Avila S.V., Souza V., Gaut B.S., Eguiarte L.E. Timing and rate of speciation in agave (agavaceae) Proc Natl Acad Sci U S A. 2006;103(24):9124–9129. doi: 10.1073/pnas.0603312103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Koch M.A., Haubold B., Mitchell-Olds T. Comparative evolutionary analysis of chalcone synthase and alcohol dehydrogenase loci in Arabidopsis, Arabis, and related genera (Brassicaceae) Mol Biol Evol. 2000;17(10):1483–1498. doi: 10.1093/oxfordjournals.molbev.a026248. [DOI] [PubMed] [Google Scholar]
  • 30.Levin D.A. The timetable for allopolyploidy in flowering plants. Ann Bot. 2013;112(7):1201–1208. doi: 10.1093/aob/mct194. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Singer T., Yordan C., Martienssen R.A. Robertson's mutator transposons in A. thaliana are regulated by the chromatin-remodeling gene decrease in DNA methylation (DDM1) Genes Dev. 2001;15(5):591–602. doi: 10.1101/gad.193701. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.de la Luz Gutierrez-Nava M., Warren C.A., Leon P., Walbot V. Transcriptionally active MuDR, the regulatory element of the mutator transposable element family of zea mays, is present in some accessions of the Mexican land race Zapalote chico. Genetics. 1998;149(1):329–346. doi: 10.1093/genetics/149.1.329. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Pace J.K., Gilbert C., Clark M.S., Feschotte C. Repeated horizontal transfer of a DNA transposon in mammals and other tetrapods. Proc Natl Acad Sci U S A. 2008;105(44):17023–17028. doi: 10.1073/pnas.0806548105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Gao D. Identification of an active mutator-like element (MULE) in rice (Oryza sativa) Mol Genet Genom. 2012;287(3):261–271. [Google Scholar]
  • 35.Cresse A.D., Hulbert S.H., Brown W.E., Lucas J.R., Bennetzen J.L. Mu1-related transposable elements of maize preferentially insert into low copy number DNA. Genetics. 1995;140(1):315–324. doi: 10.1093/genetics/140.1.315. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Lisch D. Mutator transposons. Trends Plant Sci. 2002;7(11):498–504. doi: 10.1016/s1360-1385(02)02347-6. [DOI] [PubMed] [Google Scholar]
  • 37.Lisch D., Girard L., Donlin M., Freeling M. Functional analysis of deletion derivatives of the maize transposon MuDR delineates roles for the MURA and MURB proteins. Genetics. 1999;151(1):331–341. doi: 10.1093/genetics/151.1.331. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Benito M.I., Walbot V. Characterization of the maize mutator transposable element MURA transposase as a DNA-Binding protein. Mol Cell Biol. 1997;17(9):5165–5175. doi: 10.1128/mcb.17.9.5165. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Cosby R.L., Chang N.C., Feschotte C. Host-transposon interactions: conflict, cooperation, and cooption. Genes Dev. 2019;33(17–18):1098–1116. doi: 10.1101/gad.327312.119. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Munasinghe M., Springer N., Brandvain Y. Critical role of insertion preference for invasion trajectory of transposons. Evolution. 2023;77(10):2173–2185. doi: 10.1093/evolut/qpad128. [DOI] [PubMed] [Google Scholar]
  • 41.Hunter C.T., McCarty D.R., Koch K.E. Independent evolution of transposase and TIRs facilitated by recombination between mutator transposons from divergent clades in maize. Proc Natl Acad Sci U S A. 2023;120(31) [Google Scholar]
  • 42.Gao D., Caspersen A.M., Hu G., Bockelman H.E., Chen X. A novel mutator-like transposable elements with unusual structure and recent transpositions in barley (Hordeum vulgare) Front Plant Sci. 2022;13 [Google Scholar]
  • 43.Dupeyron M., Singh K.S., Bass C., Hayward A. Evolution of mutator transposable elements across eukaryotic diversity. Mobile DNA. 2019;10:12. doi: 10.1186/s13100-019-0153-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Hisanaga T., Romani F., Wu S., Kowar T., Wu Y., Lintermann R., et al. The polycomb repressive complex 2 deposits H3K27me3 and represses transposable elements in a broad range of eukaryotes. Curr Biol. 2023;33(20):4367–43680 e9. doi: 10.1016/j.cub.2023.08.073. [DOI] [PubMed] [Google Scholar]
  • 45.Hure V., Piron-Prunier F., Yehouessi T., Vitte C., Kornienko A.E., Adam G., et al. Alternative silencing states of transposable elements in Arabidopsis associated with H3K27me3. Genome Biol. 2025;26(1):11. doi: 10.1186/s13059-024-03466-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Pélissier T., Jarry L., Olivier M., Dajoux G., Pouch-Pélissier M.-N., Courtois C., et al. Plant mobile domain protein–DNA motif modules counteract Polycomb silencing to stabilize gene expression. Nat Plants. 2025 doi: 10.1038/s41477-025-02127-1. [DOI] [Google Scholar]
  • 47.Ono A., Kim S.H., Walbot V. Subcellular localization of MURA and MURB proteins encoded by the maize MuDR transposon. Plant Mol Biol. 2002;50(4–5):599–611. doi: 10.1023/a:1019970206057. [DOI] [PubMed] [Google Scholar]
  • 48.Donlin M.J., Lisch D., Freeling M. Tissue-specific accumulation of MURB, a protein encoded by MuDR, the autonomous regulator of the mutator transposable element family. Plant Cell. 1995;7(12):1989–2000. doi: 10.1105/tpc.7.12.1989. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Raizada M.N., Benito M.I., Walbot V. The MuDR transposon terminal inverted repeat contains a complex plant promoter directing distinct somatic and germinal programs. Plant J. 2001;25(1):79–91. doi: 10.1046/j.1365-313x.2001.00939.x. [DOI] [PubMed] [Google Scholar]
  • 50.Borredá C., Leduque B., Colot V., Quadrana L. Transposable element products, functions, and regulatory networks in arabidopsis. bioRxiv [Preprint] 2024. Available from: 10.1101/2024.04.02.587720. [DOI]
  • 51.Velanis C.N., Perera P., Thomson B., de Leau E., Liang S.C., Hartwig B., et al. The domesticated transposase ALP2 mediates formation of a novel polycomb protein complex by direct interaction with MSI1, a core subunit of polycomb repressive complex 2 (PRC2) PLoS Genet. 2020;16(5) [Google Scholar]
  • 52.Crescente J.M., Zavallo D., Helguera M., Vanzetti L.S. MITE tracker: an accurate approach to identify miniature inverted-repeat transposable elements in large genomes. BMC Bioinf. 2018;19(1):348. [Google Scholar]
  • 53.Potter S.C., Luciani A., Eddy S.R., Park Y., Lopez R., Finn R.D. HMMER web server: 2018 update. Nucleic Acids Res. 2018;46(W1):W200–W204. doi: 10.1093/nar/gky448. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54.Miele V., Penel S., Duret L. Ultra-fast sequence clustering from similarity networks with SiLiX. BMC Bioinf. 2011;12:116. [Google Scholar]
  • 55.Solovyev V., Kosarev P., Seledsov I., Vorobyev D. Automatic annotation of eukaryotic genes, pseudogenes and promoters. Genome Biol. 2006;7(Suppl 1):S10 1–2. Suppl 1. [Google Scholar]
  • 56.de Vienne D.M. Lifemap: exploring the entire tree of life. PLoS Biol. 2016;14(12) [Google Scholar]
  • 57.Katoh K., Misawa K., Kuma K., Miyata T. MAFFT: a novel method for rapid multiple sequence alignment based on fast fourier transform. Nucleic Acids Res. 2002;30(14):3059–3066. doi: 10.1093/nar/gkf436. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 58.Nei M., Gojobori T. Simple methods for estimating the numbers of synonymous and nonsynonymous nucleotide substitutions. Mol Biol Evol. 1986;3(5):418–426. doi: 10.1093/oxfordjournals.molbev.a040410. [DOI] [PubMed] [Google Scholar]
  • 59.Kimura M. A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J Mol Evol. 1980;16(2):111–120. doi: 10.1007/BF01731581. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Multimedia component 1
mmc1.xlsx (7.1MB, xlsx)
Multimedia component 2
mmc2.pdf (262KB, pdf)

Data Availability Statement

Information regarding genome assemblies and active autonomous TE clusters is available in the online Supplementary Data.

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