The genome of Populus trichocarpa was retrieved from Phytozome v12.1. The amino acid sequences of A. thaliana WRKY proteins were downloaded from TAIR (https://www.arabidopsis.org/). Using these known A. thaliana WRKY proteins as query sequences, a BLASTP search (v2.2.28; E-value ≤ 1×e^-10) was performed against the P. trichocarpa genome database. Concurrently, WRKY genes were identified using HMMER 3.0 with the WRKY domain (PF03106) Hidden Markov model (HMM, E-value ≤ 1×e^-10) from the Pfam database (http://pfam.xfam.org). We took the intersection of the results from these two approaches. Sequences lacking intact WRKYGQK motifs or zinc finger domains (C₂HC/C₂H₂ type) were filtered out via multiple sequence alignment and validated using SMART 9.0 (http://smart.embl-heidelberg.de/) and NCBI Conserved Domain Database (CDD).

The physicochemical properties and subcellular localization of P. trichocarpa WRKY proteins were predicted using ProtParam (https://web.expasy.org/protparam/) and WoLF PSORT (https://wolfpsort.hgc.jp/). The amino acid sequences of WRKY genes from P. trichocarpa and A. thaliana were selected for phylogenetic relationship analysis. A phylogenetic tree was constructed using the Maximum Likelihood (ML) method in MEGA12 software (with 1000 bootstrap replicates), and optimization was conducted using the iTOL platform (https://itol.embl.de/). Conserved motifs were identified using MEME (http://meme-suite.org/), with the maximum number of motifs set to 10, the minimum motif width to 6, and the maximum motif width to 50. Functional annotation was performed with InterProScan (https://www.ebi.ac.uk/interpro/). Secondary structures were predicted using SOPMA (https://npsa-prabi.ibcp.fr/cgi-bin/npsa_automat.pl?page=npsa_sopma.html), tertiary structures were generated using SWISS-MODEL (https://swissmodel.expasy.org/), and topological heterogeneity models were analyzed using Protter (http://wlab.ethz.ch/protter/start/). Tandem and segmental duplication events within the P. trichocarpa WRKY gene family were analyzed using MCScanX (Python) with default parameters (BLASTP E-value< 1×e^-10, match score >50). Chromosomal localization of WRKY genes was visualized using TBtools v2.210. Collinearity relationships between P. trichocarpa and A. thaliana, Solanum lycopersicum (GCF_000188115.5), Eucalyptus grandis (GCF_016545825.1), and Glycine max were performed using the Dual Synteny Plotter (TBtools v2.210). Promoter regions of WRKY genes (2000 bp upstream of the start codon) were extracted. Cis-regulatory elements within these promoter regions were predicted using the PlantCARE database (https://bioinformatics.psb.ugent.be/webtools/plantcare/html/). Ka/Ks ratios for duplicated gene pairs were calculated using KaKs_Calculator (v2.0). Genomic data for all species were obtained from Phytozome v12.1 (https://phytozome-next.jgi.doe.gov/) and the NCBI (https://www.ncbi.nlm.nih.gov/).

Wild-type (WT) Populus davidiana × P. alba var. Pyramidalis (Pdpap) provided by the Department of Forest Protection, Northeast Forestry University (Harbin, Heilongjiang, China) was used for genetic transformation. Leaves from 1- to 2-month-old Pdpap seedlings were placed on differentiation medium (0.5 g·L^-1 6-BA; 0.1 g·L^-1 NAA; 2.47 g·L^-1 1/2MS basal medium; 20 g·L^-1 sucrose; pH 6.0) and cultured for 2 months, then transferred to rooting medium (2.47 g·L^-1 1/2MS basal medium; 0.01 mg·L^-1 NAA; 20 g·L^-1 sucrose; pH 6.0). After an additional 2 months, plants were acclimatized in a soil culture room. Both tissue culture and soil culture rooms were maintained at 22°C with a 16/8 h (light/dark) photoperiod and 70% relative humidity. Soil-cultured seedlings of Pdpap with consistent growth status were selected for subsequent treatments.

B. dothidea was isolated from canker-infected poplar stems collected at the demonstration forestry base of Northeast Forestry University. Pathogen purification was performed using tissue isolation on PDA medium (200 g·L^-1 potato infusion; 15 g·L^-1 agar; 20 g·L^-1 glucose; pH 7.0). The strain was deposited in the China General Microbiological Culture Collection Center (CGMCC 3.27823). B. dothidea was cultured on PDA for 7 d until sporulation. For inoculation, a 6-mm epidermal incision was made on the stem of Pdpap seedlings 2–3 cm above the soil surface. A 6-mm mycelial plug of B. dothidea was placed on the incision, while a sterile PDA plug served as the control. Growth conditions for Pdpap were identical to those described in Section 2.3. Leaf samples were collected at 0, 6, 12, 24, and 48 h post-inoculation to analyze the expression pattern of PdpapWRKY11 during infection. Three biological replicates were included for each time point. Additionally, root, stem, and leaf samples were collected at 0 h for spatial expression profiling, with three biological replicates per tissue.

Total RNA was extracted from Pdpap using the RNAprep Pure Plant Plus Kit (Tiangen, Beijing, China), and cDNA synthesized with TransScript First-Strand cDNA Synthesis SuperMix (TransGen, Beijing). PdpapWRKY11 was amplified with gene-specific primers under the following PCR conditions: initial denaturation at 95°C for 5 min; 35 cycles of 94°Cfor 30 s (denaturation), 58°C for 30 s (annealing), 72°C for 1 min (extension); followed by a final extension at 72°C for 10 min, gel-purified (EasyPure^® Quick Gel Extraction Kit), cloned into DH5α (Vazyme Biotech Co., Ltd., Nanjing, China), and sequenced (Primer sequences are listed in Supplementary Table 1). Evolutionary relationships were analyzed via BLAST on NCBI, and phylogenetic trees were built using MEGA X (neighbor-joining method). Amino acid sequence alignment was performed with DNAMAN 9.

Total RNA was extracted from samples obtained in Section 2.4 using the method described in Section 2.5. Subsequently, cDNA was synthesized to analyze the expression pattern of PdpapWRKY11 during B dothidea infection and its spatial expression profile. qPCR was performed using SYBR Green PCR Master Mix (Vazyme Biotech Co., Ltd., Nanjing, China) and gene-specific primers, with 100 ng of cDNA template per amplification reaction. Three independent biological replicates were analyzed per group, and PdpapEF1-α served as the reference gene for qPCR normalization. Relative expression levels were calculated using the 2^−ΔΔCT method (primer sequences are listed in Supplementary Table 1).

The fusion construct 35S::PdpapWRKY11-GFP (containing the PdpapWRKY11 coding sequence and GFP) and control 35S-GFP were transiently transformed into A. thaliana protoplasts using established protocols. Localization was observed under a Nikon C2-ER confocal microscope (Nikon Instruments Inc., Tokyo, Japan) with excitation at 488 nm and emission detection at 510 nm.

The PdpapWRKY11 gene obtained in Section 2.5 was cloned into the pBI121 vector (SpeI/XmaI sites) under the 35S promoter. The recombinant plasmid (pBI121-PdpapWRKY11) was transformed into Agrobacterium tumefaciens EHA105 (Verdure Biotech Co., Ltd., Nanjing, China). Bacterial cultures (OD₆₀₀ = 0.5–1.0) were used for leaf disc transformation of WT poplar. Transformed explants were cultured for 2 months followed by 1 month soil acclimatization under conditions described in Section 2.3. Genomic DNA was extracted from 3-month-old transformant lines using a Super Plant Genomic DNA DP360 Kit (Tiangen Biotech, Beijing) and subjected to PCR verification with gene-specific and vector primers (PdpapWRKY11-F/pBI121-R, PdpapWRKY11-R/pBI121-F). Controls included: recombinant plasmid (positive control), WT plants (negative control), and ddH₂O (no-template control). The PCR program followed the protocol described in Section 2.5. qPCR was further performed to confirm transcription levels (primers in Supplementary Table 1).

To evaluate the impact of PdpapWRKY11 overexpression on B. dothidea resistance, two-month-old WT and transformant lines (OE4, OE7, OE10) were inoculated with B. dothidea at three months of age using the protocol described in Section 2.4. Plants were maintained under growth conditions identical to Section 2.3 for 20 d. Fresh weight and root length were measured at 0, 5, 10, 15, and 20 d, with three biological replicates per group. Baseline values (0 d WT) were normalized to 1(relative value normalization). Concurrently, disease indices were statistically analyzed for both WT and transformant lines. Based on the disease progression observed in the plants, lesions were classified into five severity grades: Grade 0 (healthy/no lesions), Grade 1 (lesion width< 1/3 of stem circumference), Grade 2 (lesion width 1/3–1/2 of stem circumference), Grade 3 (lesion width 1/2–3/4 of stem circumference), and Grade 4 (lesion width > 3/4 of stem circumference or plant mortality). The disease index was calculated using the formula (Jilin Provincial Administration for Market Regulation (JQTS), 2013):

For each enzyme assay, 0.05 g of leaf tissue was pulverized in liquid nitrogen and homogenized in 0.1 mol/L phosphate buffer (pH=7.0). The homogenate was centrifuged at 10,000 × g for 10 min at 4 °C, and the supernatant was used for measurements. Peroxidase (POD) activity was quantified by colorimetric assay (Li et al., 2013), superoxide dismutase (SOD) by the hydroxylamine method (Zhao et al., 2013), malondialdehyde (MDA) content by the thiobarbituric acid (TBA) method (Liu et al., 2010), and hydrogen peroxide (H₂O₂) levels by colorimetric assay (Liu et al., 2013), using commercial kits from Nanjing Jiancheng Bioengineering Institute (Nanjing, China) strictly adhering to the manufacturer’s protocols. The corresponding catalog numbers were A084-3-1, A001-1-2, A003-3-1, and A064-1-1.

Leaf samples (0.05 g) were collected at 0, 6, 12, 24, and 48 h post-B. dothidea inoculation, pulverized in liquid nitrogen, and extracted with 2 mL of 60% ethanol under agitation at 60 °C for 2 h (Xu J, et al., 2021). After centrifugation at 10,000 × g (room temperature, 10 min), the supernatant was collected for flavonoid quantification using a colorimetric assay kit (Nanjing Jiancheng Bioengineering Institute, China; Cat. No. A142-1-1). Stem samples collected at 0, 5, 10, 15, and 20 d post-inoculation were oven-dried at 80 °C, pulverized, sieved through a 30-mesh filter, and 5 mg aliquots were used for lignin content determination via the acetyl bromide method (Solarbio Science & Technology Co., Ltd., Beijing, China; Cat. No. BC4200) with three biological replicates (Hu et al., 2022).

Leaf samples (0.05 g) were collected at 0, 6, 12, 24, and 48 h post-B. dothidea inoculation for each enzyme assay. Phenylalanine ammonia-lyase (PAL) activity (Liu et al., 2005) was quantified using a colorimetric assay kit (Nanjing Jiancheng Bioengineering Institute, China; Cat. No. A137-1-1), while cinnamyl alcohol dehydrogenase (CAD) activity (De Ascensao et al., 2000) was determined via an NADH-dependent activity assay kit (Boxbio Science & Technology Co.,Ltd., Beijing, China; Cat. No. AKSU053M). RNA from these samples was used for qRT-PCR analysis of PdpapPAL and PdpapCAD expression, with PdpapEF1-α as reference genes (primer sequences listed in Supplementary Table 1).

The qRT-PCR and physiological data were analyzed using two-way ANOVA and one-way ANOVA (SPSS 24.0, IBM), followed by Student’ s t-test for pairwise comparisons, with a P-value< 0.05 considered statistically significant.

In P. trichocarpa, 102 WRKYs were systematically identified and designated as PtrWRKYs (numbered according to their genomic positions in the P. trichocarpa genome; Supplementary Table 2). To elucidate evolutionary relationships among PtrWRKYs, a ML phylogenetic tree was constructed using amino acid sequences from P. trichocarpa and A. thaliana, enabling subfamily classification (Figure 1). According to the phylogenetic topology and classification criteria for A. thaliana WRKYs, the PtrWRKYs were stratified into seven distinct clades, with each clade color-coded for visualization (Robatzek et al., 2000). Subfamily IIc exhibited the largest membership (25 PtrWRKYs), whereas subfamily IIa contained the fewest members (5 PtrWRKYs). Subfamilies I, IIb, IId, IIe, and III comprised 23, 9, 13, 14, and 13 PtrWRKYs, respectively. Notably, homologs of AtWRKYs were randomly distributed across all subfamilies.

The physicochemical properties of PtrWRKYs exhibited substantial variation across different subfamilies, while members within the same group demonstrated highly consistent characteristics (Supplementary Table 3). The PtrWRKY proteins had an average length of 371 amino acids (range: 135–732 aa) with corresponding molecular weights averaging 41.00 kDa (range: 15.54–78.91 kDa). Theoretical isoelectric points (pI) spanned from 5.08 to 9.86, and aliphatic indices varied between 31.85 and 82.00. Instability index analysis revealed values of 37.83–70.28, with 92% of proteins classified as unstable (>40) and only a minority as stable (<40). All PtrWRKY members exhibited hydrophilic properties, as evidenced by their grand average of hydropathicity (GRAVY) scores between -1.34 and -0.36.

To investigate the structural characteristics and functional evolution of the PtrWRKYs, this study revealed the co-evolutionary patterns between molecular architecture and functional modules through integrated analyses of Motif Pattern, Conserved domain, and Gene structure (Supplementary Figure 1). MEME (Multiple Expectation Maximization for Motif Elicitation) identified 10 conserved motifs (Motif1-Motif10) in the PtrWRKYs, annotated via Pfam. Motif1 contained the canonical WRKYGQK domain, while Motif2/4/6 formed the DNA-binding framework, with Motif4 harboring a zinc finger. Motif5 exhibited intrinsic disorder, Motif8 was classified into the PRK13922 super family, and the remaining motifs lacked functional annotations. Phylogenetic clustering revealed that each PtrWRKY gene carries 2–8 motifs, with evolutionarily related members displaying highly conserved motif combinations. The ubiquitous presence of Motif2 across all PtrWRKYs, subfamily-specific distributions (e.g., Motif5 enriched in subfamilies I/IIb/IIc, Motif8 predominant in IIa/IIb, Motif9 exclusive to IIa/IIb/IIe, and Motif10 unique to subfamily I).

Conserved Domain analysis further uncovered structural evolutionary signatures: while subfamily I retains dual WRKY domains, other subfamilies possess a single WRKY domain. Acquired domains, including Plant_zn_clust in IId (zinc binding), FlaE domain in WRKY89 (flavonoid synthesis), Nucleoporin_FG2 superfamily in WRKY100 (transport through nuclear pore), and ODAM superfamily (enamel formation) in WRKY30, indicate domain-mediated functional specialization. Gene structure analysis revealed conserved genomic architectures among subfamily members, with intron numbers ranging from 1 to 6 (predominantly 2-4) across different subgroups. The results demonstrated that subfamilies I, IIa, IIb, IIc, IId, IIe, and III contained 2-6, 2-4, 3-6, 1-3, 2-3, 1-3, and 2 introns, respectively. The findings collectively demonstrate that the functional diversity of the WRKY family arises from the synergistic interplay of motif combinatorial patterns, domain variations, and exon-intron structural divergence.

Secondary structure analysis (Supplementary Table 4) revealed that PtrWRKY family proteins predominantly consist of α-helices (13.78%, 49 amino acids on average), extended strands (6.72%, 24 amino acids), and random coils (79.50%, 295 amino acids), with no β-turns detected. Structural heterogeneity was observed among members: PtrWRKY29 exhibited the highest α-helix proportion (29.47%), PtrWRKY1 displayed the most extended strands (10.37%), and PtrWRKY64 showed exceptionally high random coil content (87.79%), suggesting potential conformational flexibility. The tertiary structure modeling (Supplementary Table 5) was performed using high-similarity templates from the AlphaFold Database, with sequence identities ranging from 70.13% to 100.00% (average 89.80%). All models were validated as high-confidence structures.

The topological heterogeneity model analysis revealed that no transmembrane helices were identified in any PtrWRKY proteins (Supplementary Table 6). Specifically, 13 PtrWRKYs lacked N-glycosylation sites, whereas 89 PtrWRKYs contained 0–12 predicted N-glycosylation sites. None of the PtrWRKYs exhibited detectable signal peptide sequences. Subcellular localization predictions demonstrated that the majority of PtrWRKYs were nuclear-localized (Supplementary Table 7). Notably, PtrWRKY99 was predicted to reside in the cytoplasm, PtrWRKY18 and PtrWRKY76 in peroxisomes, and PtrWRKY90 in chloroplasts.

The chromosomal distribution of PtrWRKYs in poplar exhibits pronounced non-randomness and spatial heterogeneity (Supplementary Figure 2). These genes are distributed across 18 out of 19 chromosomes, with complete absence on Chr09, suggesting potential inhibitory epigenetic regulatory mechanisms suppressing WRKY gene evolution or the absence of adaptive evolutionary pressures on this chromosome. Chr01 and Chr02 display significant gene enrichment, each harboring 12 PtrWRKY members (accounting for 11.76% of the family). Notably, gene distribution density shows no correlation with chromosomal length: the shorter chromosome 6 (approximately 50.2 Mb) contains 9 genes, while the longer chromosome 16 (72.8 Mb) accommodates only 4 members. Remaining chromosomes demonstrate graded distribution patterns: chromosomes 5, 12, and 17 each carry 6 genes; chromosomes 3, 4, and 14 contain 5 each; chromosomes 7, 8, 10, and 19 host 3 each; chromosomes 13 and 16 possess 2 members.

A substantial proportion of plant gene duplication arises from interchromosomal replication. To elucidate the evolutionary origins and expansion mechanisms of the PtrWRKYs in poplar, this study integrates intra- and inter-species collinearity analyses to reveal its duplication patterns. Intra-species collinearity analysis using MCScanX identified extensive segmental duplication events among PtrWRKYs, with 58 core genes forming 95 homologous duplication pairs (Figure 2A). These pairs, interconnected by red lines in the synteny map, are distributed across 18 chromosomes harboring PtrWRKYs and exhibit interchromosomal interaction features, indicating that interchromosomal recombination dominates gene family expansion.

Collinearity-based inference suggests that one gene in each duplicated pair originated from segmental duplication of the other. The Nonsynonymous to synonymous substitution ratio (Ka/Ks), a metric for evolutionary rate and selective pressure, revealed that all duplication pairs exhibit Ka/Ks values below 1 (mean = 0.24, range: 0.07–0.53), strongly supporting purifying selection.

To assess cross-species conservation of WRKY TFs, we constructed comparative synteny maps between P. trichocarpa and four representative dicot species (A. thaliana, Solanum lycopersicum, Eucalyptus grandis, and Glycine max) (Figure 2B). Results demonstrate broad evolutionary conservation between PtrWRKYs and their dicot homologs, with the highest collinearity observed in G. max (soybean, 400 homologous events), followed by E. grandis (149 events), S. lycopersicum (142 events), and A. thaliana (118 events). In summary, both interchromosomal segmental duplication events and functional conservation of the gene family influence the functional differentiation of genes in PtrWRKYs.

To better understand the biological processes involved in PtrWRKYs, this study systematically revealed subfamily-specific regulatory networks by analyzing 2,000 bp cis-acting elements in the promoter regions of PtrWRKYs (Supplementary Figure 3). A total of 27 functional cis-acting elements were identified, with each gene harboring 19–66 elements (mean = 34), which were classified into four core regulatory modules: (1) stress response module, including hypoxia/anaerobic, drought, pathogen, and pest stress elements; (2) growth and development module, such as meristem expression, endosperm expression, root-specific elements, and differentiation of palisade mesophyll cells; (3) hormone response module, encompassing ABA, methyl jasmonate (JA signaling), auxin, and salicylic acid (SA); (4) photoperiod regulation module containing light-responsive elements and circadian control. Notably, all PtrWRKYs carried these four element categories but exhibited significant functional divergence among subfamilies: in biotic stress response elements, subfamilies IIa/IIb/IIc were enriched with as-1 (biotic stress response), while IIe/IId preferentially contained WRE3 (wound response), and IId/III subfamilies specifically accumulated WUN-motif (damage response). Hormone signaling elements (MYC, ARE) were highly enriched in I/III/IIe subfamilies, whereas growth and development-related elements exhibited the lowest overall proportion. These findings demonstrate that the functional expression of the PtrWRKYs genes is regulated by cis-acting elements associated with light, phytohormones, stress, and plant development.

Based on phylogenetic and functional modular analyses of the WRKY gene family in P. trichocarpa, we found that multiple subfamilies (e.g., IId and III) were significantly enriched with stress-responsive cis-elements, suggesting their potential central roles in stress resistance regulation in the Populus genus.

To validate the biological functions of WRKY TFs in poplar-pathogen interactions, this study employed Pdpap as the experimental system. This hybrid poplar exhibits strong innate disease resistance against the canker pathogen B. dothidea and possesses well-established genetic transformation protocols. By integrating evolutionary theoretical findings from P. trichocarpa with functional validation in this high-resistance hybrid system, this study advanced the elucidation of plant disease resistance mechanisms.

Our research group previously conducted transcriptome sequencing on Pdpap infected with B. dothidea at five critical timepoints (0, 6, 12, 24, and 48 h) to investigate the resistance mechanisms of Pdpap against the pathogen. Based on RNA-seq analysis using thresholds of |log₂FC| > 2 and FDR< 0.01, four genes were persistently up-regulated and three genes consistently showed down-regulated trends across all treatment groups compared to 0 h Pdpap seedlings (Supplementary Figure 4, Supplementary Table 8). Among the four up-regulated genes, only PdpapWRKY11 belongs to Subgroup IId of P. trichocarpa (previously identified as a stress-responsive subfamily). Among these, PdpapWRKY11 (homologous to the stress-responsive subfamily IId of P. trichocarpa) was thus selected for subsequent experiments. The PdpapWRKY11 gene was cloned from PdPap. The sequence contains a 1,017 bp open reading frame (ORF) encoding 338 amino acids. Phylogenetic analysis revealed high homology between PdpapWRKY11 and WRKY11 sequences from P. alba, P. trichocarpa, P. nigra, and P. euphratica (Figure 3A). Multiple sequence alignment of PaWRKY11, PtWRKY11, PnWRKY11, PeWRKY11, and PdpapWRKY11 showed 98.66% identity, with the highest similarity to P. alba. Conserved domain analysis confirmed the presence of WRKY and Plant_zn_clust domains (Figure 3B).

To determine the intracellular localization of PdpapWRKY11, the pBWA(V)HS-GFP vector was digested with EcoRV and ligated to PdpapWRKY11 via homologous recombination. GFP fluorescence, visualized under a confocal laser scanning microscope, confirmed nuclear localization of the pBWA(V)HS-PdpapWRKY11-GFP fusion protein in transfected Arabidopsis protoplasts (Figure 3C).

Tissue-specific expression analysis revealed that PdpapWRKY11 transcripts were most abundant in roots (2.02-fold higher than leaves) and least in leaves, with intermediate levels in stems (1.35-fold higher than leaves)(Figure 3D). Under B. dothidea infection, PdpapWRKY11 expression was significantly induced, peaking at 12 h (2.86-fold higher than 0 h) and declining to 2.15-fold by 24 h (Figure 3E). All infected groups showed markedly higher expression than the 0 h control, confirming B. dothidea responsive activation of PdpapWRKY11.

Ten PdpapWRKY11 transformant Pdpap lines were generated via Agrobacterium-mediated transformation. PCR with gene-specific and vector primers (PdpapWRKY11-F/pBI121-R, PdpapWRKY11-R/pBI121-F) confirmed transgene integration in all lines, with bands matching the positive control (pBI121-PdpapWRKY11) and negative controls (water) or no amplification in WT (Supplementary Figure 5). qRT-PCR identified OE4, OE7, and OE10 as high-expressing lines, while other lines showed suboptimal expression. RNA-based PCR validation confirmed overexpression in all 10 lines, with OE4, OE7, and OE10 selected for subsequent studies (Figure 4A).

To investigate the role of PdpapWRKY11 in Pdpap’s response to B. dothidea, transformant Pdpap lines (OE4, OE7, OE10) and WT seedlings were inoculated with the pathogen. As shown in Figure 4B, WT seedlings exhibited severe leaf abscission and stem lesions after inoculation, while OE lines maintained healthy growth (the pathological manifestations are shown in Supplementary Figure 6), indicating that PdpapWRKY11 overexpression enhances resistance to B. dothidea. Statistical analysis of disease indices revealed that the WT exhibited a significantly higher disease index (83%) compared to the transformant lines (20%).

Fresh weight analysis (Figure 4C) revealed no significant changes in OE4 and OE7 during T0-T4, whereas WT showed significant reductions at all time points. OE10 displayed significant fresh weight changes only during T0-T1 and T3-T4. Root length remained stable across all lines (Figure 4D), but OE plants exhibited superior growth under pathogen stress. These results demonstrate that PdpapWRKY11 overexpression mitigates the negative impact of B. dothidea on seedling growth.

The antioxidant role of PdpapWRKY11 was analyzed by measuring antioxidant enzyme activities (POD, T-SOD) and oxidative damage markers (H₂O₂, MDA). Under B. dothidea stress, all lines showed an initial increase followed by a decline in POD and T-SOD activities. During early infection (T0-T1), OE lines exhibited significantly higher POD (16.92–18.64% increase) and T-SOD (10.73–19.46% increase) activities than WT (Supplementary Figures 7A, B).

H₂O₂ levels exhibited stage-specific regulation patterns (Supplementary Figure 7C). Under B. dothidea stress, all lines showed a dynamic “initial decrease followed by increase” in H₂O₂ content. At T0, H₂O₂ levels in OE4 and OE7 showed significantly lower levels. By T1, WT and OE10 accumulated more H₂O₂ than OE4 and OE7. At T2, the transformant Pdpap lines transiently surpassed WT in H₂O₂ accumulation. During T3-T4, H₂O₂ levels in OE lines were significantly lower than in WT. These results indicate that PdpapWRKY11 dynamically regulates reactive oxygen species (ROS) homeostasis to balance disease resistance responses.

MDA levels followed a similar “initial decrease followed by increase” trend as H₂O₂ (Supplementary Figure 7D). During early infection (T0), MDA content in OE lines was 20.27-96.72% lower than in WT. However, by T3, OE7 showed significantly higher MDA levels than WT, potentially linked to stage-specific lipid peroxidation regulation. These findings demonstrate that PdpapWRKY11 enhances systemic resistance against canker disease by temporally regulating antioxidant enzyme activities and oxidative damage markers.

To investigate the metabolic regulatory role of PdpapWRKY11 in Pdpap’s defense against B. dothidea, lignin and flavonoid accumulation patterns were systematically analyzed in transformant Pdpap lines (OE4, OE7, OE10) and WT plants. Under pathogen stress, all lines exhibited a dynamic “initial increase followed by decrease” in lignin and flavonoid content. During early infection (0–10 d), OE lines accumulated significantly more lignin than WT. At 10 d post-inoculation, lignin levels in OE lines were 30.20–37.43% higher than in WT (P < 0.01). In later stages (10–20 d), lignin content declined in all lines (Figure 5A). Flavonoid levels showed no significant differences between OE and WT during T0-T1. However, during T2-T4, WT exhibited 29.26–40.00% higher flavonoid accumulation than OE lines (P < 0.01) (Figure 5B). These results suggest that PdpapWRKY11 preferentially enhances lignin biosynthesis, which plays a critical role in defense against B. dothidea.

To elucidate how PdpapWRKY11 regulates lignin biosynthesis under B. dothidea stress, the activities of key phenylpropanoid pathway enzymes (PAL and CAD) and related gene expression were systematically measured. Under pathogen stress, all lines showed an “initial increase followed by decrease” in PAL and CAD activities. During T0-T1, PAL activity in OE lines was significantly higher than in WT, with a 30.04–39.58% increase at T1. CAD activity remained elevated in OE lines compared to WT during T0-T2 (Figures 5C, D).

qRT-PCR results confirmed significant upregulation of PdpapPAL and PdpapCAD in OE lines, consistent with enzyme activity trends (Figures 5E, F). These findings indicate that PdpapWRKY11 may activate PdpapPAL and PdpapCAD to enhance lignin biosynthesis, thereby strengthening pathogen defense.