Abstract
Chloroplasts are key players in photosynthesis and immunity against microbial pathogens. However, the precise and timely regulatory mechanisms governing the control of photosynthesis-associated nuclear genes (PhANGs) expression in plant immunity remain largely unknown. Here we report that TaPIR1, a Pst-induced RING-finger E3 ubiquitin ligase, negatively regulates Pst resistance by specifically interacting with TaHRP1, an atypical transcription factor histidine-rich protein. TaPIR1 ubiquitinates the lysine residues K131 and K136 in TaHRP1 to regulate its stability. TaHRP1 directly binds to the TaHRP1-binding site elements within the PhANGs promoter to activate their transcription via the histidine-rich domain of TaHRP1. PhANGs expression induces the production of chloroplast-derived ROS. Although knocking out TaHRP1 reduces Pst resistance, TaHRP1 overexpression contributes to photosynthesis, and chloroplast-derived ROS production, and improves disease resistance. TaPIR1 expression inhibits the downstream activation of TaHRP1 and TaHRP1-induced ROS accumulation in chloroplasts. Overall, we show that the TaPIR1-mediated ubiquitination and degradation of TaHRP1 alters PhANGs expression to disrupt chloroplast function, thereby increasing plant susceptibility to Pst.
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Introduction
Plants are continuously exposed to several potential pathogens. In response, they have evolved a refined, two-branched defense system to protect themselves against such disease-causing agents1,2,3. The first layer of defense involves multiple plasma membrane-localized pattern recognition receptors that can detect pathogen-associated molecular patterns (PAMPs) such as chitin and flagellin (Flg22), which elicit various immune responses, including callose deposition, reactive oxygen species (ROS) accumulation, and defense-related gene expression4. To achieve successful invasion, pathogens deliver various effectors into host cells to suppress their basal immune response by interfering with host immune factors, such as receptors and downstream signaling factors. In addition, plants employ specific disease resistance genes (R genes) to detect pathogen-secreted effectors. The strong interaction between effectors and R proteins remarkably induces disease resistance that rivals the specificity and range of the mammalian adaptive immune system5. Notably, the production of immune signals such as ROS and nitric oxide (NO) induces salicylic acid (SA) synthesis at infection sites. Increased SA levels further elicit plant defense responses in a self-amplifying feedback loop, ultimately leading to a hypersensitive response (HR)—a type of cell necrosis that occurs at infection sites to restrict pathogen spread6,7.
In plants, chloroplasts play key roles in immunity as major sites for the production of carbohydrates and the generation of various defense signals, such as SA, Ca2+, and NO signals. In chloroplasts, isochorismate synthase catalyzes the conversion of chorismate into isochorismate, which is further converted into SA8,9. A chloroplast-localized protein, calcium-sensing receptor (CAS), plays a role in the formation of chloroplast-derived Ca2+ transients and regulates SA biosynthesis to control plant innate immunity10. SA acts as a key regulator not only in PAMPs-triggered immunity and effectors-triggered immunity (ETI) but also in systemic acquired resistance, which refers to broad-spectrum and systemic resistance to secondary infections. Importantly, when plant tissues encounter biotic or abiotic stress, chloroplasts can reprogram and integrate secondary metabolite synthesis, phytohormone crosstalk, and communication with other organelles via retrograde signaling11,12. In addition, chloroplasts are crucial contributors to redox homeostasis, including ROS generation. The ROS in chloroplasts includes hydroxyl radicals (OH•), hydrogen peroxide (H2O2), and singlet-excited oxygen (1O2)13. Excess light absorption to the maximum capacity of the photosynthetic electron transport chain (PETC) for CO2 assimilation induces the production of 1O2 and H2O2 from photosystem I (PSI) and PSII14. Excessive ROS accumulation limits pathogen colonization at infection sites. Moreover, ROS serves as retrograde signaling molecules that induce nuclear gene expression. Several studies have provided insights into chloroplast-mediated plant immunity against microbial pathogens. In Arabidopsis, the reduction of PsbS, a subunit of photosystem II, contributed to energy dissipation by protecting the photosystems from damage caused by excess light and enhanced photosynthesis-induced ROS accumulation after the perception of PAMP15. In wheat, the chloroplast-targeted kinase WKS1 encoded by the rust resistance gene Yr36 interacts with and phosphorylates PsbO, a PSII oxygen-evolving enhancer protein 1, to disrupt the photosystem and confer resistance to Pst16. The stable silencing of TaISP, which encodes a subunit of the Cyt b6/f complex, inhibits photosynthesis and chloroplast-derived ROS accumulation, resulting in the reduced resistance of plants to avirulent Pst17. Although numerous studies on chloroplast function have reported the role of chloroplast-targeting proteins in plant immunity, the precise and timely regulatory mechanisms of transcription in controlling the expression of chloroplast-associated nuclear genes in wheat remain largely unknown.
Ubiquitination, an essential post-translational regulatory process, involves the enzymatic attachment of a chain of ubiquitin molecules to target proteins. This modification via a megacomplex protease, 26S proteasome, marks the target proteins for degradation18,19. Ubiquitination is achieved through the sequential activities of the ubiquitin-activating enzyme E1, ubiquitin-conjugating enzyme E2, and ubiquitin ligase E320. In plants, ubiquitination E3 ligases can be subdivided into three classes, namely, U-box, homologous to E6-associated protein C-terminus, and really interesting new gene (RING)-type E3 ligases21. Several studies have reported the role of ubiquitination in various cellular processes, including hormone responses, development, and immunity. In rice, microtubule-associated E3 ligase ubiquitinates serine hydroxymethyltransferase and promotes its degradation to induce broad-spectrum resistance22. In addition, two E3 ubiquitin ligases, RIP1 and APIP6, fine-tuned the Ca2+ sensor ROD1 to inhibit ROS scavenging by reducing the catalase activity of CatB, ultimately conferring resistance to multiple pathogens23,24. By contrast, the rice scaffold protein OsCUL3a of the E3 ubiquitin ligase complex interacts with and degrades OsNPR1 via 26S proteasome, thus negatively affecting cell death and plant immunity25. However, compared with the ubiquitination processes in rice, the role of ubiquitin ligase E3 in wheat immunity and its mode of substrate recognition remains unclear.
Herein, we identified a RING-finger ubiquitin E3 ligase, TaPIR1, as a susceptibility factor that plays a negative role in inducing Pst resistance in wheat. The TaPIR1 knockout in hexaploid wheat induced broad-spectrum resistance to the Pst races CYR31 and CYR34. TaPIR1 physically interacted with and ubiquitinated an atypical transcription factor histidine-rich protein, TaHRP1, which directly regulates photosynthesis-associated nuclear genes (PhANGs) expression. Although the loss of TaHRP1 drastically reduced Pst resistance in wheat, its overexpression contributed to chloroplast-derived ROS accumulation and improvement in disease resistance. The TaPIR1-induced binding and modification of TaHRP1 to suppress PhANGs expression may affect the production of chloroplast-derived ROS, as well as photosynthesis. Overall, these findings indicate that TaPIR1-mediated ubiquitination and degradation of TaHRP1 act as a switch to disrupt chloroplast function and ROS accumulation, increasing plant susceptibility to Pst.
Results
TaPIR1 is a negative defense regulator against Pst in wheat
Among the differentially expressed genes in Pst-infected wheat leaves26, the expression of TaPIR1−4A (Puccinia striiformis-induced RING-finger protein 1), a member of the E3 ubiquitin-protein ligase family, was induced at 24 h post-inoculation (hpi), as described in the quantitative reverse transcription polymerase chain reaction (qRT-PCR) results (Supplementary Fig. 1a). Despite 99% nucleotide sequence similarity among the three copies of TaPIR1 located on chromosomes 4AS, 4BS, and 4DS, the TaPIR1-4A and TaPIR1-4B transcripts exhibited significant induction incompatible interactions, approximately by threefold, at 24 hpi; however, the TaPIR1-4D transcripts remained undetected (Supplementary Fig. 1a). To understand the role of TaPIR1 in wheat–Pst interaction, we first employed barley stripe mosaic virus (BSMV)-induced gene silencing to knock down TaPIR1 expression. At 10 days after BSMV inoculation, photobleaching was observed in Phytoene desaturase (TaPDS)-silenced plants, which served as a positive control. Compared with the control plants, the TaPIR1-silenced wheat plants exhibited fewer urediniospore pustules and a typical HR when wheat plants expressing the BSMV constructs were infected with Pst CYR34 at 14 days post-infection (dpi) (Supplementary Fig. 1b). The TaPIR1 transcript exhibited a 60–70% reduction at 24 hpi and 48 hpi in plants expressing the BSMV construct of TaPIR1 (Supplementary Fig. 1c). The TaPIR1-silenced plants exhibited lower Pst biomass than the control plants (Supplementary Fig. 1d). Furthermore, we overexpressed TaPIR1 in Fielder plants and inoculated with Pst CYR23. Compared with the control plants, the TaPIR1-expressing plants exhibited urediniospore pustules (Supplementary Fig. 1e). Consistently, the TaPIR1-overexpressing plants showed a >60–70% increase in fungal biomass compared with that of the controls (Supplementary Fig. 1f), indicating a negative role in wheat stripe rust resistance.
To further validate the role of TaPIR1 in inducing plant susceptibility to Pst, the three TaPIR1 homologs (TaPIR1-4A, TaPIR1-4B, and TaPIR1-4D) in the wheat cultivar Fielder were edited via CRISPR–Cas9 gene editing. However, only two mutant plants, which contained nucleotide deletions leading to frameshift mutations in the region targeted by Cas9 in TaPIR1-4A and TaPIR1-4B (tapir1-AB) or TaPIR1-4A alone (tapir1-A), were obtained using three pairs of primers specific to the abovementioned three TaPIR1 homologs (Supplementary Fig. 2a). When inoculated with the predominant Pst races CYR31 and CYR34, the T2 generation of tapir1-AB and tapir1-A plants exhibited a strong HR and contained a few urediniospore pustules (Fig. 1a and Supplementary Fig. 2b). Notably, compared with the wild-type (WT) plants, tapir1-AB and tapir1-A plants exhibited 50–60% reductions in fungal biomass (Fig. 1b and Supplementary Fig. 2c). The stable knockout of tapir1-AB also reduced the hyphal length and infection areas of Pst CYR34 at 24 hpi and 48 hpi (Fig. 1c, d). The tapir1-AB plants demonstrated markedly greater ROS accumulation than Fielder plants at 24 hpi and 48 hpi (Fig. 1e, f). These results indicate that TaPIR1 negatively regulates Pst resistance in wheat.
The second leaves of the seedlings of the TaPIR1 knockout plant (tapir1-AB) and Fielder were infected with Pst CYR31 and CYR34, respectively. a Pst uredinia on the second leaves of tapir1-AB and Fielder were observed at 14 dpi. b The Pst/wheat biomass ratio was measured at 120 hpi via qRT-PCR of DNA isolated from the same set of wheat plants, as shown in (a). The internal control genes TaEF and PstEF were used to normalize the DNA levels of wheat and Pst, respectively. Values are mean ± standard deviation (SD), n = 3 biologically independent samples. The P value was determined by a two-tailed unpaired Student’s t-test. c Representative images were obtained from tapir1-AB and Fielder plants inoculated with Pst CYR34 at 24 hpi or 48 hpi using CellSens Entry software. IH infectious hyphae, SV substomatal vesicle, bar = 20 μm. d Hyphal length and infection areas on the same set of wheat plants, as shown in (c), were evaluated using CellSens Entry software. Means ± SD were calculated from 30 infection sites of three independent biological repeats. The P value was determined by a two-tailed unpaired Student’s t-test. e Histological analysis of H2O2 accumulation in tapir1-AB and Fielder plants inoculated with Pst CYR34 at 24 hpi and 48 hpi. SV substomatal vesicle, IH infectious hyphae, bar = 20 μm. f H2O2 accumulation in the same wheat samples was assessed at 24 hpi and 48 hpi using CellSens Entry software as described above. Means ± SD were calculated from 30 infection sites of three independent biological repeats. The P value was determined by a two-tailed unpaired Student’s t-test. Source data are provided as a Source Data file.
TaPIR1 interacts with and ubiquitinates TaHRP1
To verify the predicted nuclear localization sequence (NLS) in the C-terminal region, the TaPIR1:green fluorescent protein (GFP) construct driven by the CaMV35s promoter was first generated for transient infiltration in Nicotiana benthamiana. GFP signals were specifically observed in the nucleus of the resulting tobacco cells expressing TaPIR1:GFP, whereas control-free GFP was distributed throughout the cells (Supplementary Fig. 3a). Repeated experiments in wheat protoplasts revealed the specific nuclear localization of the green fluorescence of TaPIR1:GFP upon transient transfection of the TaPIR1:GFP construct (Supplementary Fig. 3b).
To further explore the molecular mechanism by which TaPIR1 influences the plant immune response, we used TaPIR1-4B as a bait to screen the yeast two-hybrid (Y2H) library of Pst-infected wheat leaves for TaPIR1-interacting proteins. In total, 45 clones were identified as 32 plant proteins (Supplementary Data 1). Owing to the nucleus localization of TaPIR1, we focused on nucleoproteins. Among the 32 interactants, all 13 nucleus-related interactants were identified and used to detect interactions with TaPIR1. Only TaHRP1, heavy metal-associated isoprenylated protein (TaHMAI), and histone H1 (TaH1) were confirmed to interact with TaPIR1 by the Y2H system. The yeast cells expressing pBD:TaPIR1–pAD:TaHRP1, pBD:TaPIR1–pAD:TaH1, and pBD:TaPIR1–pAD:TaHMAI grew on SD-Ade plates and exhibited galactosidase activity (Fig. 2a and Supplementary Data 1).
a Detection of TaPIR1–TaHRP1 interaction using the yeast two-hybrid assay. Yeast transformants with marked constructs were grown on SD medium lacking LW (-Leu/-Trp) and those with LacZ activities were grown on SD-LWHA (-Leu/-Trp/-His/-Ade) supplemented with X-α-gal. b Detection of TaPIR1–TaHRP1 interaction in the nucleus of N. benthamiana leaves expressing the corresponding constructs based on the BIFC assay. A NLS fused with a red fluorescent protein (RFP) was used as a nucleus marker. Bar = 20 µm. Similar results are obtained from two independent biological experiments. c Verification of TaPIR1–TaHRP1 interaction using GST pull-down assay. Western blots of the recombinant protein mixture and proteins eluted from GST beads were detected with anti-His or anti-GST antibodies. Similar results are obtained from three independent biological experiments. d Confirmation of the TaPIR1–TaHRP1 interaction via Co-IP assays. Total proteins from N. benthamiana leaves expressing the labeled constructs and proteins copurified from GFP beads were detected via western blotting with anti-GFP and anti-HA antibodies. Similar results are obtained from two independent biological experiments. e Ubiquitination of TaHRP1 by TaPIR1 in vitro. Ubiquitinated TaHRP1, GST-tagged TaHRP1, and His-tagged TaPIR1 were separated and detected via western blotting with anti-ubiquitin, anti-GST, and anti-His antibodies, respectively. Similar results are obtained from two independent biological experiments. f Confirmation of TaHRP1 ubiquitination by TaPIR1 in protoplasts prepared from tapir1-AB, TaPIR1-overexpression, tapir1-AB-expressing TaPIR1 and Fielder plants. Ubiquitinated proteins copurified from the total proteins were detected with an anti-ubiquitin antibody. Similar results are obtained from two independent biological experiments. Source data are provided as a Source Data file.
To further analyze the physical interaction between TaPIR1 and its interactants, they were fused with the C- and N-terminal portions of yellow fluorescent protein (YFP), respectively, using the bimolecular fluorescence complementation (BIFC) assay. The fused products were transformed into Agrobacterium tumefaciens and then infiltrated into tobacco leaves. The fluorescence signals of the TaPIR1–TaHRP1 interaction were specifically observed in the nucleus (Fig. 2b). By contrast, fluorescence signals remained undetected when TaPIR1:cYFP–TaH1:nYFP and TaPIR1:cYFP–TaHMAI:nYFP were transiently expressed (Fig. 2b). Furthermore, the in vitro pull-down assay indicated that glutathione S-transferase (GST):TaHRP1 can effectively pull down TaPIR1:His compared with the GST control, suggesting a physical interaction between TaPIR1 and TaHRP1 (Fig. 2c). In addition, we performed the coimmunoprecipitation (Co-IP) assay in tobacco leaves cotransformed with TaPIR1:GFP/TaHRP1:HA, as well as the total protein extracted from tobacco leaves and mixed with GFP-trap beads. HRP1:HA was coimmunoprecipitated by TaPIR1:GFP but not by GFP alone, indicating that TaPIR1 interacts with TaHRP1 in vivo (Fig. 2d). To identify the TaHRP1-interacting region, we artificially truncated the disordered region in TaPIR1 predicted via DISOPRED 3 and generated bait constructs of TaPIR1 mutants (Supplementary Fig. 4a). However, all TaPIR1-truncated mutants failed to interact with TaHRP1 in yeast (Supplementary Fig. 4b). Furthermore, TaPIR1 interacted with TdHRP1 derived from Triticum durum but not with HvHRP1 derived from Hordeum vulgare, although their protein sequences were 80% identical to the TaHRP1 sequence (Supplementary Fig. 4c–e). Overall, these findings confirm that TaPIR1 physically interacts with TaHRP1 in the nucleus.
To analyze E3 ligase activity in TaPIR1, we first expressed and purified His-tagged TaPIR1:His derived from Escherichia coli (Supplementary Fig. 3c) and performed an in vitro ubiquitination assay. When TaPIR1:His was incubated with purified E1, E2, and ubiquitin, a protein ladder of polyubiquitinated TaPIR1:His was detected using anti-ubiquitin antibodies, indicating that TaPIR1 possesses E3 ligase activity and can mediate self-ubiquitination in vitro (Supplementary Fig. 3d). However, TaPIR1M, in which His-60 and His-235 were replaced by alanine in the RING structure, lost E3 ligase activity in vivo (Supplementary Fig. 3e). We further analyzed whether TaHRP1 can be used as a ubiquitination substrate for TaPIR1. In the presence of TaPIR1:His, E1, E2, and TaHRP1:GST, a strong protein ladder of polyubiquitinated TaHRP1:GST was detected (Fig. 2e). However, in the absence of E1, E2, TaPIR1:His (E3) or TaHRP1, no TaHRP1 polyubiquitination was observed (Fig. 2e). To further determine whether TaPIR1 contributes to TaHRP1 ubiquitination in vivo, immunoprecipitation assay was performed. As expected, more ubiquitinated TaHRP1 proteins were detected in the protoplasts prepared from TaPIR1OE or tapir1-AB-expressing TaPIR1 compared with Fielder plants; however, ubiquitinated TaHRP1 proteins rarely appeared in the protoplasts prepared from tapir1-AB (Fig. 2f). Consistent with this result, an in vivo ubiquitination assay revealed that TaHRP1 is ubiquitinated in N. benthamiana co-expressing TaPIR1. However, TaPIR1M was not observed to ubiquitinate TaHRP1 in N. benthamiana (Supplementary Fig. 5a). Collectively, these results demonstrate that TaPIR1 is a functional E3 Ub ligase that can ubiquitinate the TaHRP1 protein.
TaPIR1 promotes TaHRP1 degradation
Considering that TaPIR1 directly interacts with TaHRP1 and ubiquitinates it both in vitro and in vivo, we determined whether the E3 ligase TaPIR1 affects TaHRP1 protein turnover. TaHRP1 stability was evaluated based on cell-free degradation analysis of the total protein extracts derived from the loss-of-function mutants tapir1-AB, Fielder plants, and TaPIR1-overexpressing plants. Compared with GST control, TaHRP1:GST accelerated degradation in coincubation mixtures from Fielder plants but were relatively stable in coincubation mixtures from tapir1-AB compared with Fielder plants extracts (Fig. 3a and Supplementary Fig. 5b). In contrast to the Fielder plants extracts, incubation with extracts from TaPIR1-overexpressing plants significantly increased the degradation rate of TaHRP1:GST (Fig. 3a). In all coincubation mixtures, the presence of MG132 inhibited TaHRP1:GST degradation (Fig. 3a). In addition, TaHRP1:GFP exhibited a lower amount of protein in TaHRP1-overexpressing plants infected with Pst CYR34 compared with that in uninfected plants (Supplementary Fig. 2g). In addition, more TaHRP1 proteins were detected in the protoplasts prepared from tapir1-AB compared with Fielder plants; however, the TaHRP1 protein abundance was reduced in the protoplasts prepared from tapir1-AB-expressing TaPIR1 or TaPIR1OE plants (Supplementary Fig. 3f). Consistent with this result, an in vivo degradation assay revealed that TaPIR1 promoted TaHRP1 degradation in wheat protoplasts co-expressing TaPIR1 but TaPIR1M could not (Supplementary Fig. 5c). We obtained similar results in N. benthamiana, wherein TaHRP1 protein levels were affected by TaPIR1, but TaPIR1M did not have the same effect (Fig. 3b). Based on these results, we concluded that TaPIR1 ubiquitinates TaHRP1, leading to its degradation by the 26S proteasome.
a Cell-free degradation assay of TaHRP1 in coincubation mixtures from Fielder (WT), tapir1-AB, and TaPIR1OE plants. TaHRP1:GST derivatives expressed and purified in E. coli were incubated for different times at 28 °C with protein extracts from Fielder (WT), tapir1-AB, and TaPIR1-overexpressing plant leaves. The relative intensities (R-value) of each band were evaluated using ImageJ software and are shown above the bands. The bands in the left electrophoresis line are reference bands (set to 1). b In vivo degradation of TaHRP1 in N. benthamiana leaves transiently expressing TaPIR1. Protein levels were detected via western blotting with anti-GFP antibody in the presence of 50 μM MG132 or an equal volume of DMSO (control). The relative intensities (R-value) of each band are shown above the bands, and the bands in the left electrophoresis line are used as a reference (set to 1). Actin was used as the loading control. c Cell-free degradation assay of mutants of TaHRP1 detected in coincubation mixtures from Fielder (WT) incubated for different times at 28 °C. The relative intensities (R-value) of each band are shown above the bands, and the bands in the left electrophoresis line are reference bands (set to 1). Actin was used as the loading control. d Mutants of TaHRP1 in in vivo accumulation assay. Different mutants of TaHRP1 were coexpressed with TaPIR1 in N. benthamiana leaves. The relative intensities (R-value) of each band are shown above the bands, and the bands in the right electrophoresis line are used as a reference (set to 1). Actin was used as the loading control. Source data are provided as a Source Data file.
TaHRP1 degradation depends on its lysine residues K131 and K136
We further analyzed the potential ubiquitination sites of TaHRP1 and found that six lysine residues were present in the C-terminal region of TaHRP1. K131 and K136 residues of TaHRP1 protein were predicted as the most likely ubiquitination sites (Supplementary Fig. 5d). To confirm whether TaPIR1 can catalyze the ubiquitination of these sites, we expressed and purified TaHRP1 mutants and performed an in vitro ubiquitination assay. When the lysine residues K131 and K136 were mutated into the arginine residues R131 and R136, respectively, the extent of ubiquitination of TaHRP1K131R and TaHRP1K136R was not significantly different compared with that of TaHRP1 (Fig. 2e). In contrast, TaHRP1K131,136R ubiquitination remained inconspicuous (Fig. 2e). We also evaluated the ubiquitination of three mutants of TaHRP1 in vivo. Western blotting analysis of proteins eluted from GFP-trap beads revealed some amount of ubiquitinated TaHRP1K131R and TaHRP1K136R but not of ubiquitinated TaHRP1K131,136R upon their transient expression in tobacco (Supplementary Fig. 5e). To exclude the possibility of a relationship between three TaHRP1 mutants and TaPIR1 may influence ubiquitination of TaHRP1, these interactions were analyzed through the Y2H assays. Similar to TaHRP1, three mutants were found to interact with TaPIR1 (Supplementary Fig. 4f). These results demonstrate that TaPIR1 can ubiquitinate the K131 and K136 residues of the TaHRP1 protein.
To further determine the key lysine sites responsible for TaHRP1 stability, we conducted a cell-free degradation assay to detect the stability of three mutants of TaHRP1 by incubating total protein extracts from tapir1-AB, WT, and TaPIR1-overexpressing plants, respectively. When incubated with WT or TaPIR1-overexpressing plant extracts, the degradation rates of TaHRP1K136R:GST and TaHRP1K131R:GST increased but remained relatively stable in coincubation mixtures from tapir1-AB plant (Fig. 3c and Supplementary Fig. 6). In contrast, TaHRP1K131,136R remained relatively stable in all coincubation mixtures (Supplementary Fig. 6). Repeated experiments revealed TaPIR1, but not TaPIR1M, significantly promoted TaHRP1K131R and TaHRP1K136R protein degradation when three mutants of TaHRP1 and TaPIR1 or TaPIR1M was transiently co-expressed in N. benthamiana (Fig. 3d). No changes were observed in TaHRP1K131,136R protein abundance (Fig. 3d). Overall, these findings consistently demonstrate that TaPIR1 ubiquitinates K131 and K136 sites of TaHRP1 protein, thereby mediating its degradation.
TaHRP1 is a positive regulator of wheat resistance to Pst
To determine the role of TaHRP1 in Pst infection, we first conducted CRISPR–Cas9-mediated gene editing to fully edit three TaHRP1 homologs in the wheat cultivar Fielder. Subsequently, we identified a tahrp1 line in which three TaHRP1 homologs contained deletions leading to frameshift mutations (Supplementary Fig. 2d). Upon infection with avirulent Pst CYR23, a typical resistance response with HR was observed in Fielder plants; in contrast, tahrp1 plants presented with evident urediniospore pustules (Fig. 4a). TaPR1 and TaPR2 transcription was reduced 2–4-fold at 0 hpi and 24 hpi in tahrp1 plants compared with that in Fielder plants (Supplementary Fig. 7a, b). Consistently, tahrp1 plants showed >50–60% increase in fungal biomass relative to the controls (Fig. 4b). Moreover, histochemical examination of Pst revealed longer hyphae and larger infection areas in the tahrp1 plant than in the control (Fig. 4c). In repeated experiments, the hyphal length and infection areas of Pst CYR23 increased compared with the control (Fig. 4d, e). These results confirm that the loss of TaHRP1 markedly reduces resistance to avirulent Pst.
a The second leaves of the seedlings of the T2 generation of tahrp1 and Fielder plants were inoculated with Pst CYR23. Leaves infected with Pst urediniospores were observed at 14 dpi. b The biomass ratio (Pst/wheat) was measured using DNA isolated from the second leaves inoculated with Pst at 120 hpi. Wheat TaEF and rust PstEF were used to normalize the DNA level. Values are mean ± SD, n = 3 biologically independent samples. Data was analyzed using an unpaired two-tailed Student’s t-test. c Pst growth and development in tahrp1 and Fielder plants inoculated with Pst CYR23 at 24 hpi and 48 hpi. Representative images were observed using CellSens Entry software. IH infectious hyphae, SV substomatal vesicle, bar = 20 μm. d, e Hyphal length and infection areas were analyzed using CellSens Entry software at 24 hpi and 48 hpi. Means ± SD were calculated from 30 infection sites of three independent biological repeats. The P value was determined by a two-tailed unpaired Student’s t-test. f Pst uredinia on the second leaves of TaHRP1OE and Fielder's plants inoculated with Pst CYR34 were obtained at 14 dpi. g Fungal biomass at 14 dpi with Pst CYR34 was determined via qPCR. TaEF and PstEF were used as internal control genes to measure wheat and Pst DNA with reference to the standard curves. Values are mean ± SD, n = 3 biologically independent samples. The P value was determined by a two-tailed unpaired Student’s t-test. h Relative expression of the marked defense-related genes in TaHRP1OE plants at 0 hpi and 24 hpi with Pst CYR34. Transcript levels were confirmed via qRT-PCR and normalized with TaEF. Values are mean ± SD, n = 3 biologically independent samples. The P value was determined by a two-tailed unpaired Student’s t-test. Source data are provided as a Source Data file.
To further determine the role of TaHRP1 in wheat stripe rust resistance, we generated transgenic wheat lines overexpressing TaHRP1 (TaHRP1OE). The transcript levels of TaHRP1 in TaHRP1OE#L3 and #L5 plants increased threefold and fourfold, respectively (Supplementary Fig. 2e). YFP-tagged TaHRP1 was also detected via western blotting with an antibody against GFP, indicating the successful expression of TaHRP1 in TaHRP1OE#L3 and #L5 plants (Supplementary Fig. 2f). When inoculated with virulent Pst CYR34, Fielder plants developed numerous urediniospore pustules and showed no obvious HR at 14 dpi, whereas TaHRP1OE#L3 and L5 plants developed sporadic urediniospore pustules and showed HR (Fig. 4f). TaHRP1OE#L3 and L5 plants showed 40–50% reduction in Pst biomass at 120 hpi relative to WT plants (Fig. 4g). In addition, we measured the expression of two defense-related genes in transgenic wheat lines overexpressing TaHRP1. TaPR1 and TaPR2 expression increased 2–3-fold at 0 hpi and 24 hpi in the TaHRP1OE#L3 and L5 plants compared with that in Fielder plants (Fig. 4h). Taken together, TaHRP1 may function as a positive regulator of wheat stripe rust resistance.
TaHRP1 acts as a transcription factor directly targeting PhANGs
As TaHRP1 is homologous to the forkhead box protein B2 that binds to a cis-acting element27, we hypothesized that TaHRP1 acts as a transcription factor in the nucleus. To test this hypothesis, we first confirmed the predicted NLS in the C-terminal region. Upon transient expression of TaHRP1:GFP in N. benthamiana, GFP signals were predominantly observed in the nucleus, with only a portion of TaHRP1:GFP accumulating in the cytoplasm (Supplementary Fig. 8a, b). Similar results were confirmed in the nuclear extracts of N. benthamiana leaves transiently expressing TaHRP1:GFP. A high accumulation of TaHRP1:GFP was detected on the western blots of proteins isolated from the plant nucleus (Supplementary Fig. 8c).
Next, we assessed the self-transactivation activity of TaHRP1 in yeast. Yeast cells expressing pBD:TaHRP1 were cultured on SD-His plates supplemented with 25 mM 3-AT and exhibited galactosidase activity (Supplementary Fig. 8d). To identify the self-transactivation activity region, two different fragments of TaHRP1 were amplified and tested in yeast cells. The self-transactivation activity of TaHRP1 was eliminated after the C-terminal region was deleted but not after the deletion of the histidine-rich region, indicating that the C-terminal region of TaHRP1 is required for its self-transactivation activity (Supplementary Fig. 8d).
Based on the Y2H assay, the TaHRP1 was found to interact with itself, indicating its homodimer-forming ability (Supplementary Fig. 9a). When the cysteine residue C46 was mutated into an alanine residue in TaHRP1, the mutant TaHRP1C46A eliminated its interaction with TaHRP1 (Supplementary Fig. 9a). Subsequently, we performed Co-IP and split-luciferase complementation imaging assays to further confirm the TaHRP1–TaHRP1 interaction (Supplementary Fig. 9b, c). In addition, the BIFC assay revealed that the homointeraction between TaHRP1 and TaHRP1 mainly occurred in the plant nucleus (Supplementary Fig. 9d). The purified TaHPR1 protein was then analyzed via native polyacrylamide gel and immunoassay to detect TaHPR1 oligomerization. In the absence of β-mercaptoethanol (β-ME), the TaHPR1 oligomers were detected but the TaHRP1C46A mutant could not undergo oligomerization (Supplementary Fig. 9e).
To further identify TaHRP1-regulated target genes, DNA affinity purification sequencing (DAP-seq) was employed to determine its genome-wide binding sites. Overall, 4.43%, 88%, and 2.74% of TaHRP1-binding peaks were distributed in promoter, intergenic, and intron regions, respectively (Supplementary Fig. 10a). We identified 27,876 and 17,839 loci, with an overlap of 5440 target peaks (Supplementary Fig. 10b). TaHRP1-binding peaks were enriched around the transcription start sites, indicating that these peaks are strongly enriched in promoter regions (Supplementary Fig. 10c). Motif enrichment analysis of the TaHRP1-binding peaks revealed that the AAGAAGAAG element (e-value = 6.3e-2067) was the most enriched motif in both DAP-seq replicates, hereafter named as HBS (Fig. 5a). KEGG analysis revealed that the potential TaHRP1-binding genes involved in photosynthesis were markedly enriched, among which four encode PSII proteins (PsbI-1A, PsbE-2D, PsbE-5D, and PsbD-4D), three encode cytochrome b6f subunits (PetA-3B, PetN-6B, and PetN-2D), four encode PSI proteins (PsaI-7D, PsaI-1A, PsaA-2B, and PsaA-3B) and one encodes ATP synthase subunit (AtpH-4A) (Fig. 5b). The heatmap revealed that, compared with the control, the DNA-binding peaks of the abovementioned PhANGs were enriched by TaHRP1 in the two DAP-seq replicates (Fig. 5c). Consistently, visual assessment of the binding peaks of PhANGs confirmed that the notable conspicuous peaks were present in their promoter regions (Fig. 5d and Supplementary Fig. 10d). These results indicate that TaHRP1 may regulate the expression of PhANGs.
a The conserved binding motifs of TaHRP1 protein were predicted using MEME-ChIP. The core sequence of “AAGAAGAAG” was substantially enriched among the TaHRP1 binding regions (e-value = 2.2e-2067) and was named HBS. b The top 10 enriched KEGG terms of the TaHRP1 direct targets. The q value was defined based on Fisher’s test. c Heatmap showing fold enrichment of 12 PhANGs directly regulated by TaHRP1. Rep1 and Rep2, are two replicates of TaHRP1 relative to the control. The fold enrichment was relative to the control, and the scale bar shows log2 (FPKM). d IGV browser view of TaHRP1 binding to TaPsbE-2D. Tracks show data from the control, as well as Rep_1 and Rep_2, two replicates of TaHRP1. e Binding of recombinant TaHRP1:GST protein to the TaPsbE-2D promoter was analyzed via EMSA. The HBS element is shown in purple in the promoter region. The TaHRP1:GST fusion protein concentrations were 2 μg and 1 μg, and the GST protein concentration was 2 μg (negative control). Similar results are obtained from two independent biological experiments. f Confirmation of the binding affinity of TaHRP1 and the 60-bp TaPetN promoter containing the HBS element using the isothermal titration calorimetry (ITC) assay. Fitted integrated ITC data curve of the 60-bp TaPetN promoter with TaHRP1, TaHRP1H/A:GST, and control GST. The calculated binding kinetic dissociation constant (Kd values ± errors) for the promoter with TaHRP1 was 0.14 ± 0.08 μM and that with TaHRP1H/A:GST was 0.36 ± 0.15 mM. TaHRP1H/A is a mutant of TaHRP1 in which all histidine residues in the N-terminal region are mutated into alanine residues. Source data are provided as a Source Data file.
TaHRP1 regulates PhANG expression
DAP-seq revealed that the promoter region exhibited considerable enrichment of PhANGs. To validate the reliability of our DAP-seq results, the promoter fragment of PsbE-2D containing the HBS element was isolated and incubated with the TaHRP1:GST protein using the electrophoretic mobility shift (EMSA) assays. Evidently, a specific band appeared only when TaHRP1:GST was included. With an increase in TaHRP1:GST concentration, a band indicative of binding increased in abundance, whereas the band was completely absent in the HBS element-mutated probe control (Fig. 5e and Supplementary Fig. 11a). However, all histidine residues in the histidine-rich domain in the N-terminal region of TaHRP1 mutated into alanine (TaHRP1H/A), eliminating its interaction with the HBS DNA probe (Supplementary Fig. 11b). We further examined the thermodynamics of TaHRP1–HBS interaction using isothermal titration calorimetry (ITC). When the HBS element was injected into cell pools containing GST and TaHRP1H/A:GST proteins, the heat exchange volumes were small and could not fit any binding model, indicating that there was no specific binding between the HBS element and GST or TaHRP1H/A:GST (Supplementary Fig. 11c). In contrast, considerable heat changes were observed upon injecting the HBS element into cells containing TaHRP1:GST protein (Supplementary Fig. 11c). The binding constant (Kd) values were determined to be 0.14 ± 0.08 μM for TaHRP1:GST and 0.36 ± 0.15 mM for TaHRP1H/A:GST, suggesting that the histidine-rich domain in the N-terminal region of TaHRP1 is essential for its interaction with HBS (Fig. 5f).
To investigate the mechanism through which TaHRP1 regulates PhANGs expression, we subsequently performed a dual-luciferase reporter assay to determine the effect of TaHRP1 on the transcription of 12 PhANGs. The results revealed that the expression of four PSII-related genes (PsbI-1A, PsbD-4D, PsbE-5D, and PsbE-2D), three ETC-related genes (PetA-3B, PetN-2D, and PetN-6B), and one AtpH-4A gene was significantly transactivated by TaHRP1, with the expression of four PSI-related genes (PsaI-7D, PsaI-1A, PsaA-3B, and PsaA-2B) remaining unaffected compared with that of the control (Fig. 6a). To assess the impact of TaPIR1 on TaHRP1-mediated transcriptional regulation of PhANGs, we used TaHRP1 as an effector in the dual-luciferase reporter assay of N. benthamiana leaves expressing TaPIR1:GFP and TaPIR1M:GFP. Transient expression of TaPIR1:GFP, but not TaPIR1M:GFP, suppressed the downstream activation of TaHRP1 (Fig. 6b). These conclusions were further supported by qRT-PCR results revealing the increased expression of four PSII-related genes and three ETC-related genes in tapir1-AB transgenic wheat compared with that in the controls, whereas reversed expression pattern was observed in Fielder plants expressing TaPIR1 (Supplementary Fig. 12a, b). In addition, the transcripts of PSII-related genes and ETC-related genes exhibited significant induction in incompatible interactions, approximately by 2–3-fold (Supplementary Fig. 12c). Collectively, these findings further indicate that TaPIR1 counters the role of TaHRP1 in regulating PhANGs transcription.
a Dual-luciferase reporter assay showed that the transcriptional activities of PhANGs were activated by TaHRP1. The corresponding reporter and effector constructs were simultaneously transformed into N. benthamiana leaves, and the enzyme activities of firefly luciferase (LUC) and Renilla luciferase (REN) were detected at 48 hpi. Data are presented as the mean ± SD of three biological replicates. The P value was determined by a two-tailed unpaired Student’s t-test. b Suppression of TaHRP1-activated PhANGs expression in N. benthamiana leaves transiently expressing TaPIR1. Data are presented as the mean ± SD of three biological replicates. The P value was determined by a two-tailed unpaired Student’s t-test for multiple comparisons. c Assays for electron transport rate (ETR), maximum dark-adapted quantum efficiency (Fv/Fm), and nonphotochemical quenching (NPQ) in N. benthamiana leaves infiltrated with Agrobacterium expressing TaHRP1, TaHRP1H/A, or GFP. Data are presented as the mean ± SD of three biological replicates. The P value was determined by a two-tailed unpaired Student’s t-test. d Suppression of TaHRP1-induced ROS in chloroplasts by TaPIR1. Tobacco leaves infiltrated with Agrobacterium expressing TaHRP1 and TaPIR1, TaHRP1H/A, TaPIR1 alone (control), or TaHRP1 treated with H2DCF-DA were observed for DCF signals. Bar = 20 μm. Similar results are obtained from three independent biological experiments. e N. benthamiana leaves treated with H2DCF-DA were observed via confocal microscopy for DCF signals at 48 hpi with Agrobacterium strains expressing PhANGs. Bar = 20 μm. Similar results are obtained from three independent biological experiments. Source data are provided as a Source Data file.
TaHRP1 contributes to chloroplast-derived ROS homeostasis
Considering the transcriptional activation of TaHRP1 to PhANGs, we speculated that the photoinhibited PSII system influences electron transfer efficiency and ROS accumulation. To test this hypothesis, we measured the effect of TaHRP1 on photosynthesis by overexpressing this protein in N. benthamiana and wheat plants. TaHRP1-overexpressing tobacco leaves caused a rapid increase in the maximum dark-adapted quantum efficiency (Fv/Fm) and electron transport rate (ETR) at 24 hpi and 36 hpi compared with the control (Fig. 6c). Moreover, a decrease in nonphotochemical quenching (NPQ) was observed at 24 hpi and 36 hpi in TaHRP1-overexpressing plants (Fig. 6c). In TaHRP1-overexpression wheat plants, the ETR was increased compared with the ETR in Fileder plants (Supplementary Fig. 7c). These results indicate that TaHRP1 plays a significant role in modulating photosynthesis.
As ROS are unavoidable byproducts of photosynthesis and are mainly generated in chloroplasts, we measured chloroplast-derived ROS accumulation using the probe 2′7′-dichlorodihydrofluorescein diacetate (H2DCF-DA) in tobacco leaves and wheat protoplasts overexpressing TaHRP1H/A, TaHRP1, and TaPIR1. Notably, TaHRP1 induced a high accumulation of ROS in chloroplasts but TaHRP1H/A or TaPIR1 did not exert the same effect (Fig. 6d and Supplementary Fig. 13a). Importantly, more ROS was accumulated in wheat protoplasts of TaHRP1OE and tapir1-AB plants, whereas no ROS accumulation was observed in wheat protoplasts of WT, tahrp1, or TaPIR1-overexressing plants (Supplementary Fig. 13b). In addition, we measured ROS production in TaHRP1OE plants inoculated with Pst CYR34, which revealed that H2O2 accumulation increased at 24 hpi and 48 hpi (Supplementary Fig. 13c, d). Furthermore, the influence of TaPIR1 on TaHRP1-induced ROS production was detected. Transient coexpression of TaHRP1 and TaPIR1 in tobacco gradually reduced TaHRP1-induced ROS production (Fig. 6d). To verify that ROS production was induced by PhANGs expression, we also examined ROS production in N. benthamiana leaves transiently expressing PhANGs. The DCF signal was strongly enhanced in leaves transiently expressing PetN-6B, PetA-3B, PsbE-2D, PsbI-1A, PsbE-5D, PsbD-4D, and AtpH-4A (Fig. 6e and Supplementary Fig. 14). No significant difference in the oxidized DCF signal was observed in the leaves expressing PsaA-3B and PsaI-7D (Fig. 6e and Supplementary Fig. 14). Therefore, TaHRP1 may contribute to photosynthesis and ROS production in chloroplasts to activate plant immunity, possibly by modulating PhANGs expression.
Discussion
The ubiquitin/proteasome pathway is a crucial protein quality control system in eukaryotes. There is substantial evidence to support that protein ubiquitination is an important component of regulatory networks controlling various plant cellular processes, including growth and development, as well as defense responses, such as APIP6 and RIP1 in rice23,29,30. However, there are limited reports of protein ubiquitination in wheat immunity responses to microbial pathogens. In this study, transcriptome sequencing of Pst-infected wheat leaves revealed that the expression of the gene encoding the E3 ligase TaPIR1 was highly induced at early infection stages. Simultaneous gene editing of TaPIR1 via CRISPR–Cas9 enhanced resistance to stripe rust in wheat, indicating that TaPIR1 is an important positive regulator of wheat susceptibility to Pst. In addition, to hinder nutrient uptake for pathogen development, fungal colony areas and hyphal growth were restricted in tapir1-AB wheat plants with an increase in ROS accumulation at infection sites. Modification of susceptibility genes has been widely regarded an effective measure to confer broad-spectrum and durable resistance in plants31,32. Mildew resistance locus O (MLO) was modified to enhance plant resistance to powdery mildew, and a 304-kbp targeted deletion in the MLO-B1 locus conferred robust powdery mildew resistance while maintaining wheat growth and yield33. The wheat kinase gene TaPsIPK1, a susceptibility gene, has been identified as an ideal target for inducing broad-spectrum resistance to rust in wheat34. Another study showed that the knockout of three homoalleles of TaWRKY19 improved the TaNOX10-mediated ROS burst and conferred high resistance to Pst35. Therefore, TaPIR1 may act as a genetic engineering target to provide durable resistance to this disease-causing pathogen.
TaPIR1 was shown to physically interact with the histidine-rich protein TaHRP1 in the plant nucleus, which has not been functionally characterized in wheat. We revealed that TaPIR1 acts as a typical RING-finger domain-containing E3 ligase to ubiquitinate the K131 and K136 residues of TaHRP1 protein and modulate its stability. Interestingly, TaHRP1 does not contain any known domains, except for a histidine-rich motif at its N-terminal region. Our results demonstrated that a histidine-rich motif is essential for binding to a cis-acting element as a transcription factor. Previous studies on Citrus unshiu have shown that the histidine-rich domains of dehydrin CuCOR15 contributed to the binding of DNA and metals, including Fe3+, Cu2+, and Zn2+ 36,37. Deletion of the histidine-rich motif of dehydrin OpsDHN1 in Opuntia streptacantha did not affect its dimer formation but suppressed its import into the nucleus38. Based on the results of this study, we speculated that TaHRP1 acts as an atypical transcription factor with an activation domain at the C-terminal region and a DNA-binding domain at the N-terminal region. TaHRP1 overexpression in wheat plants was shown to promote ROS accumulation and exert a strong positive effect on plant resistance to Pst; however, the loss of function of TaHRP1 in wheat resulted in reduced plant resistance. During pathogen invasion in plants, instantaneous ROS bursts in plant tissues can induce programmed cell death to inhibit pathogen growth and development. Therefore, we speculate that the E3 ligase TaPIR1 suppresses ROS accumulation to confer wheat susceptibility to Pst via direct ubiquitination and degradation of the positive factor TaHRP1.
In addition to supplementing photosynthesizing tissues with carbon sources and energy, chloroplasts are hubs for plant–pathogen interaction39,40. A reduction in the expression of PsbD (a PSII subunit) regulated by the transcription factor TPC13 was compromised in the ETI of Hyaloperonospora arabidopsidis41,42. Our results revealed that TaHRP1 overexpression promoted Fv/Fm and ETR and exerted a positive effect on plant photosynthesis. TaHRP1 activated PhANGs expression to encode core elements of PSII, cytochrome b6f complex, and ATP synthase43. Overexpression of the ATP synthase α-subunit gene reduced the viral load in both N. benthamiana and soybean plants. In several plant-pathogen interactions, reduced photosynthesis in susceptible plants promotes pathogen spread and colonization, but some resistant plants exhibit increased expression of photosynthesis-related genes in response to pathogen invasion44. Earlier studies have shown that low expression levels of the cytochrome b/f complex in transgenic tobacco can impair electron transport and reduce the CO2 assimilation rate45. The cytochrome b6/f complex and ATP synthase are considered crucial factors in the reduction of electron transport and photosynthetic capacity46. Thus, we speculate that TaHRP1 affects the photosynthetic capacity to enhance plant defense by increasing the number of core protein complexes in the photosynthetic system. Maintaining high levels of photosynthesis is important to protect PSII and PSI from damage by ROS and excess light. There is an urgent need to increase the demand for photosynthesis to provide the energy required for defense responses, which contributes to plant biosynthesis and metabolism to generate phytohormones, secondary metabolites, and defense-related proteins when organisms encounter pathogen invasion28. Furthermore, the induction of chloroplast photosynthesis genes induces hormone signaling pathways that partially contribute to local and systemic resistance to pathogens47.
Increasing evidence has shown that inhibition of photosynthesis is stock-in-trade for the suppression of plant immunity, leading to bacterial, viral, and fungal multiplication48,49,50. To suppress chloroplast function, the photosynthetic apparatus in plant immunity is usually targeted by pathogen-secreted virulence factors, such as HopN1 and Pst_1280651,52,53. The HopN1 from Pseudomonas syringae and Pst_12806 from rust fungi can interact with plant PsbQ (a protein present in the oxygen-evolving complex of PSII) and ISP, respectively, to suppress chloroplast-derived ROS accumulation51,53. The interference of pathogens with photosynthetic functions may explain why different components in the photosystem play an important role in precisely regulating chloroplast-derived ROS accumulation for plant immunity. A high ISP level can improve the photosynthetic ETR and induce ROS overproduction in PETC51. Our results showed that transient expression of TaHRP1 and its target PhANGs-related PETC promote chloroplast-derived H2O2 accumulation. However, increased levels of PsaI and PsaA, two reaction centers of PSI, failed to induce ROS in chloroplasts. In fact, the PETC and PSII of chloroplasts produce the largest pools of ROS. By contrast, the level of ROS does not change significantly in PSI reaction center54,55. Excessive ROS accumulation can accelerate chlorophyll degradation, lipid peroxidation, and photosynthesis reduction by oxidative damage to the photosystem, resulting in cell death56. Therefore, we speculate that TaHRP1 enhances photosynthetic capacity by activating PhANGs, resulting in excessive light absorption by PSII. This excess energy might be transferred from chlorophyll to molecular oxygen for ROS accumulation.
In summary, our results indicate a susceptible mechanism in which the E3 ligase TaPIR1 ubiquitinates and degrades the transcription factor TaHRP1, attenuates TaHRP1-regulated PhANGs expression, and decreases chloroplast-derived ROS accumulation (Fig. 7). During wheat–Pst interaction, when TaPIR1 expression was induced by virulent Pst, TaPIR1 directly ubiquitinated TaHRP1 to promote its degradation, which further inhibited chloroplast-derived ROS accumulation due to low expression levels of TaHRP1-regulated PhANGs, thus compromising wheat immunity. In contrast, in tapir1-AB plants, high TaHRP1 accumulation enhanced the expression of PhANGs transcripts to promote chloroplast-derived ROS accumulation and photosynthesis, which restricted infection and conferred host resistance.
The RING-finger ubiquitin E3 ligase TaPIR1 is translocated into the plant nucleus, where it interacts with and ubiquitinates the atypical transcription factor TaHRP1. TaPIR1–TaHRP1 interaction and modification promote TaHRP1 degradation and interfere with TaHRP1-induced PhANGs expression, resulting in attenuated chloroplast-derived ROS accumulation. In tapir1-AB plants, TaHRP1 accumulation activates PhANGs expression and induces ROS accumulation, which contributes to resistance to Pst in wheat.
Methods
Experimental materials and growth conditions
Wheat seedlings and N. benthamiana were grown in a greenhouse at 16 °C and 23 °C, respectively, under a light/dark photoperiod of 16 h/8 h. Wheat cultivar Fielder was used to establish TaPIR1 knockout (tapir1-A and tapir1-AB) plants, TaHRP1 knockout (tahrp1) plants, and TaHRP1 overexpression (TaHRP1OE) plants. For inoculation with Pst races CYR23, CYR31, and CYR34, wheat seedlings were grown and inoculated with urediniospores of Pst at 16 °C under a light/dark photoperiod of 16 h/8 h57. For transgenic wheat tillering and seed collection, the plant seeds from all backgrounds were grown at 25 °C with a photoperiod of 16 h of light and at 20 °C with 8 h of dark. A. tumefaciens (GV3101 and EHA105) and E. coli (DH5a and BL21) were stored at −80 °C and cultured in Luria–Bertani medium at 28 °C and 37 °C, respectively. The yeast strain (AH109) was cultured at 30 °C for the Y2H assays.
Construction of transgenic wheat lines
To perform CRISPR–Cas9 editing of TaPIR1 and TaHRP1, two candidate gRNAs targeting three copies of these genes were designed, and their editing efficiencies were determined using WheatOmics (http://202.194.139.32/)58. These gRNAs exhibiting high efficiency were introduced into the pBUE411 vector to establish the CRISPR–Cas9 vector. The A. tumefaciens strain EHA105 harboring the recombinant vector was transformed into immature embryos of the wheat cultivar Fielder via Agrobacterium-mediated transformation59. The genomic DNA of the positive transgenic seedlings was extracted for PCR detection of the Cas9 gene fragment. For constructing TaHRP1-overexpressing transgenic wheat lines, the full-length coding sequence of TaHRP1 was introduced into the pEarleyGate 104 vector with YFP at the C-terminus using Gateway technology. The recombinant vector was transformed into A. tumefaciens strain EHA105, and the transformed strains were infiltrated into the immature embryos of the wheat cultivar Fielder. The total proteins of the positive transgenic seedlings were extracted and detected via western blotting using an anti-GFP antibody (Beyotime, cat.no. AG281, 1:2000).
Subcellular localization and BIFC assays
For subcellular localization in N. benthamiana leaves, the full-length coding sequences of TaPIR1 and TaHRP1 were cloned into the pCAMBIA1302 vector and transformed into A. tumefaciens GV3101. The transformed strains with the corresponding constructs were infiltrated into N. benthamiana leaves through A. tumefaciens-mediated infiltration at a final optical density at 600 nm (OD600) of 0.6. For subcellular localization in wheat protoplasts, the full-length coding sequences of TaPIR1 and TaHRP1 were cloned into the pJIT163-GFP vector and then transfected into wheat protoplasts via PEG-mediated transfection60. The GFP signal of protoplasts was observed using a confocal microscope. For the BIFC assay, TaPIR1 and TaHRP1 were ligated with YFP in pSPYNE and pSPYCE vectors, respectively, and these constructs were transformed into A. tumefaciens GV310161. The Agrobacterium strains carrying BIFC and NLS-red fluorescent protein (RFP) recombinant vectors were infiltrated into N. benthamiana leaves at an OD600 of 0.5. Images were captured at 48 hpi via confocal microscopy using a 488 nm laser.
Yeast two-hybrid assays
The full-length coding sequence of TaPIR1 was cloned into the pGBKT7 vector as bait to screen a cDNA library of pathogen-infected wheat. The bait construct and Y2H cDNA library plasmids were cotransformed into the yeast strain AH109, which was then inoculated into an SD/-His/-Leu/-Trp/-Ade medium to screen for potential TaPIR1-interacting proteins, according to the Matchmaker Y2H system protocol (CLONTECH Laboratories). Potentially interacting proteins were screened and introduced into the pGADT7 vector as prey proteins. The bait and different prey proteins were cotransfected into the yeast strain AH109, and the diluted yeast cells were then inoculated into an SD/-Ade/-His/-Leu/-Trp medium supplemented with X-α-Gal. Images were captured after 3-day incubation at 30 °C.
Split luciferase complementation assay
The split-luciferase complementation assay was performed as previously described62. The full-length coding sequences of TaPIR1 and TaHRP1 were cloned into JW771-NLuc and JW772-CLuc vectors, respectively, and then infiltrated into N. benthamiana leaves via Agrobacterium-mediated transformation. The infiltrated plants were cultured in a greenhouse for 48 h at 23 °C, and the infiltrated leaves were smeared with d-luciferin (Coolaber, cat.no. CL6928) and incubated for 10 min under dark conditions. Images were captured using a CCD imaging system (VILBER, France).
Coimmunoprecipitation assays
For the Co-IP assay, A. tumefaciens GV3101 carrying TaHRP:HA and TaPIR1:GFP was infiltrated into N. benthamiana leaves via Agrobacterium-mediated transformation. After 48 h of infiltration, the total proteins in the infiltrated leaves were extracted using NP-40 lysis buffer (Beyotime, cat.no. P0013F) with 1 mM phenylmethanesulfonyl fluoride (Beyotime, cat.no. ST506), 1% Protease inhibitor cocktail (Beyotime, cat.no. P1005) and 50 μM MG132 (Selleck, cat.no. S2619). The extracted proteins were then incubated with GFP-trap agarose (ChromoTek, cat.no. gta-20) at 4 °C for 2 h, following the manufacturer’s instructions. The agarose beads were collected via centrifugation at 2500×g and 4 °C for 5 min and washed three times with 500 µL of phosphate buffer saline and 0.5% Tween-20. The proteins bound to GFP-trap agarose were boiled for 10 min and detected via western blotting with anti-GFP (Beyotime, cat.no. AG281, 1:2000) or anti-HA antibody (Beyotime, cat.no. AH158, 1:2000).
Nucleic and cytoplasmic extract preparation assay
TaHRP:GFP or GFP vectors were transiently expressed in N. benthamiana leaves via Agrobacterium-mediated transformation. The infiltrated leaves were ground to a fine powder using liquid nitrogen and then homogenized using Honda buffer (2.5% Dextran T40, 1.25% Ficoll 400, 10 mM MgCl2, 0.44 M sucrose, 20 mM HEPES-KOH, pH 7.4, 5 mM DTT, 1 mM PMSF, 0.5% Triton X-100, and 1% protease inhibitor cocktail) as previously described63. The extract was filtered through three layers of gauze into a 30-mL tube and then centrifuged at 3000×g and 4 °C for 10 min. The precipitates were washed three times with Honda buffer and centrifuged at 4000×g. For immunodetection, the supernatant sample and pellet resuspended in NP-40 lysis buffer were incubated with SDS loading buffer at 95 °C for 10 min.
In vitro pull-down assay
TaPIR1 and TaHRP1 fragments were cloned into pET32a and pGEX-6P-1 vectors to purify TaPIR1:His and TaHRP1:GST recombinant proteins, respectively. The tagged proteins were subjected to in vitro pull-down assay64. For affinity purification, mixtures of equal amounts of TaHRP1:GST and TaPIR1:His, as well as GST and TaPIR1:His were incubated at 4 °C for 1.5 h and mixed with glutathione resins. Recombinant protein mixtures and proteins eluted from glutathione resins were detected with anti-His (Beyotime, cat.no. AF5060, 1:2000) or anti-GST antibody (Beyotime, cat.no. AG768, 1:2000).
Ubiquitination assay
For in vivo ubiquitination assay, TaPIR1:HA or TaPIR1M:HA and TaHRP1:GFP, three mutated versions of TaHRP1:GFP (TaHRP1K131R, TaHRP1K136R, and TaHRP1K131/136R) and TaPIR1:HA were transiently expressed in N. benthamiana leaves via Agrobacterium-mediated transformation65. The infiltrated leaves were injected with 50 μM MG132 for 10 h before extracting total proteins. For the in vivo ubiquitination assay in wheat protoplasts, TaPIR1:HA and TaHRP1:GFP were co-transfected into wheat protoplasts via PEG-mediated transfection and pretreated with MG132 for 12 h before protein extraction. Proteins were extracted and incubated with GFP-trap agarose for protein enrichment. The proteins bound to GFP-trap agarose were boiled for 10 min and detected via western blotting with anti-GFP and anti-ubiquitin antibodies (HUABIO, cat.no. ET1609-21, 1:1000).
The in vitro ubiquitination assay was conducted66. The coding sequences of TaPIR1 and TaHRP1 were cloned into pET32a and pGEX-6P-1 vectors, respectively, and were purified from E. coli BL21 cells using standard protocols. Each reaction of the ubiquitination assay was performed in a 30-μL mixture containing 0.3 μg of E1 (R&D SYSTEMS, cat.no. E-305-025), 0.5 μg of E2 (R&D SYSTEMS, cat.no. E2-622-100), 2 μg of ubiquitin (R&D SYSTEMS, cat.no. U-530), 0.5 μg of TaPIR1:His, and 0.5 μg of TaHRP1:GST, mutated versions of TaHRP1:GST (TaHRP1K131R, TaHRP1K136R, and TaHRP1K131/136R) or GST, and ubiquitination buffer (50 mM Tris-HCl, PH 7.4, 2 mM ATP, 5 mM MgCl2, 2 mM DTT, 40 μM ZnSO4); this mixture was incubated at 30 °C for 3 h. For immunoblotting, the samples were incubated with an SDS loading buffer at 95 °C for 10 min. The ubiquitinated TaHRP1:GST proteins were detected with anti-ubiquitin and anti-GST antibodies, and the TaPIR1:His proteins were detected with anti-His antibodies.
Protein degradation assay
For the in vivo protein degradation assay, the TaPIR1:GFP and TaPIR1:HA recombinant vectors were transiently expressed in N. benthamiana leaves via Agrobacterium-mediated transformation. The infiltrated leaves were injected with 50 μM MG132 for 10 h before extracting total proteins using NP-40 lysis buffer. The extracted proteins were then incubated with an SDS loading buffer at 95 °C for 10 min for immunoblotting. The TaPIR1:GFP and TaPIR1:HA proteins were detected with anti-GFP and anti-HA antibodies, respectively.
For the semi-in vivo protein degradation assay, one of the four recombinant proteins—TaHRP1:GST, TaHRP1K131R:GST TaHRP1K136R:GST, TaHRP1K131,136R:GST, or GST—was incubated with the total protein extracted from Fielder, tapir1-AB, and TaPIR1-overexpressing plants in an environment supplemented with 2 mM ATP and 50 μM MG132 or DMSO at 28 °C for 0 min, 15 min, 30 min, and 60 min. TaHRP1:GST protein was detected via western blotting with an anti-GST antibody. Quantitative analysis of the immunoblots was performed using the quantity tools of ImageJ software.
DNA affinity purification and sequencing analysis
DAP-seq was performed at Bluescape Hebei Biotech following the method described in a previous study67. Briefly, genomic DNA was extracted from Fielder plants and fragmented to an average length of 200 bp using Covaris M220 (Woburn, MA, USA)68. A fragmented genomic DNA library was constructed using MICH TLX DNA-Seq Kit (Bluescape Hebei Biotech, cat.no. NGS0602). The full-length coding sequence of TaHRP1 was introduced into the pFN19K HaloTag T7 SP6 Flexi expression vector and expressed in vitro using the TNT SP6-coupled wheat germ extract system (Promega). Expressed TaHRP1 proteins were directly captured using Magne Halo Tag Beads (Promega) and incubated with the genomic DNA library; the genes were subsequently sequenced using the Illumina NavoSeq sequencing platform at Bluescape Hebei Biotech. Peak calling of the two biological duplicates was merged using MACS2 callpeak and Homer software with a q-value of <0.05, and conservative motifs were analyzed using MEME-CHIP software.
EMSA
The EMSA was performed using Light Shift Chemiluminescent EMSA Kit (Thermo Fisher Scientific, cat.no. 20158) following the manufacturer’s instructions. The promoter fragment of TaPetN containing an HBS element with a 60-bp flanking sequence upstream and downstream was biotinylated and synthesized by Sangon Biotech (Shanghai, China). The recombinant protein TaHRP1:GST, TaHRPH/A:GST, or GST at the corresponding concentration was incubated with each probe in 5× EMSA/Gel-Shift binding buffer for 30 min at room temperature. To determine the binding specificity, unlabeled or mutated biotin-labeled probes were added to the reaction mixture as the mProbe control. The products were then separated via 6% native polyacrylamide gel electrophoresis under 80 V for 1.5 h in 0.5× TBE buffer and transferred onto a positively charged nylon membrane (Millipore, cat.no. INYC00010). The binding bands on the membrane were detected using a chemiluminescent nucleic acid module. Supplementary Data 2 lists the probe sequences used in EMSA.
Transient dual-luciferase reporter assay
To evaluate the transcriptional activity of TaHRP1, promoter fragments of PhANGs containing an HBS element with a 1000-bp flanking sequence upstream and downstream were cloned into the pGreen II 0800-LUC vector, and the coding sequence of TaHRP1 was cloned into the pGreen II 62SK vector69. Supplementary Data 2 presents the primers used in this experiment. The corresponding reporter and effector constructs were simultaneously transformed into N. benthamiana leaves. After 48 h of infiltration, firefly luciferase (LUC) and Renilla luciferase (REN) activities were detected using Dual-Luciferase Reporter Assay Kit (Vazyme, cat.no. DL101) following the manufacturer’s instructions.
Detection of chlorophyll fluorescence
The chlorophyll fluorescence parameters were measured using the MINI-PAM system (Walz, Effeltrich, Germany) following the manufacturer’s instructions and a previously described protocol52. Four-week-old N. benthamiana plants were infiltrated with Agrobacterium harboring the indicated vectors and cultured under conditions of 8-h/16-h light/dark. To quantify Fv/Fm, the leaves were dark-adapted using the manufacturer’s leaf clips for 30 min at room temperature and then exposed to a saturating light pulse (5000 μmol m−2 s−1 for 0.8 s). Minimal fluorescence (Fo) and maximal fluorescence Fm were monitored with a very weak light (0.04 μmol m−2 s−1) and a transient saturated light pulse (5000 μmol m−2 s−1). Maximum quantum yield (Fv/Fm) was calculated as (Fm − Fo)/Fm. Actinic light (120 μmol m−2 s−1—the same as plant growth light intensity) was then applied for 15 min, followed by a saturating pulse to obtain maximal fluorescence (Fm´), and minimal fluorescence Fo´ after turning off the actinic light was measured. NPQ was calculated as 1 − (Fm´ − Fo´)/(Fm − Fo). Photo-irradiance response curves (LRC) were measured using different actinic light intensities (25 μmol m−2 s−1, 45 μmol m−2 s−1, 65 μmol m−2 s−1, 90 μmol m−2 s−1, 125 μmol m−2 s−1, 190 μmol m−2 s−1, 285 μmol m−2 s−1, 420 μmol m−2 s−1, 625 μmol m−2 s−1, 820 μmol m−2 s−1, 1150 μmol m−2 s−1, and 1500 μmol m−2 s−1), and each lasted for 30 s in the dark. LRC is a function of the ETR in response to PAR as shown in the following equation: ETR = PAR × ETR factor × (PPS2/PPS1+2) × Y(II), where PAR is photosynthetic active radiation, and ETR factor and (PPS2/PPS1+2) are constants with values of 0.84 and 0.5, respectively. Light intensity was monitored using WinControl 2.3 software. Measurement of 6–8 leaves was performed per replicate, and three biological replicates were examined.
BSMV-mediated gene silencing and gene overexpression
For transient silencing of TaPIR1, a specific fragment of TaPIR1 was inserted into the BSMV construct. Each BSMV construct was linearized and transcribed to RNA and then inoculated into the second leaves70. For TaPIR1 expression in wheat leaves, the full-length genes were inserted into the BSMV-VOX vector. The vector containing the tripartite viral genome and recombinant vectors was linearized and transcribed into RNA in vivo. Transcripts of each construct were mixed in a 1:1:1 ratio and then added to 200 μL of FES buffer. Seedlings treated with the virus were grown at 25–27 °C. After 10 days, the fourth leaves were inoculated with Pst urediniospores. The Pst-infected leaves were sampled at 24 hpi and 48 hpi for silencing efficiency assessment and at 120 hpi for fungal biomass assay. Leaves infected with Pst urediniospores were observed at 14 dpi.
Histochemical assay
To observe fungal hyphae, the infected leaf tissues were cleaned with ethanol and autoclaved using 1 M KOH at 121 °C for 5–6 min. Leaf fragments were carefully washed three times with 2 mL of 50 mM Tris–HCl (pH 7.4) and incubated in wheat germ agglutinin solution (Alexa-488; Thermo Fisher Scientific)71. Hyphal length and infection areas were observed using an Olympus BX-51 microscope.
For the H2O2 detection assay, the infected leaf tissues were incubated in 1.0 mg/mL DAB solution under light conditions for 4 h at 16 °C. Subsequently, the samples were treated with ethanol, as mentioned above, to remove chlorophyll, and H2O2 was observed and measured as a reddish-brown color under microscopy using the CellSens Entry program72. We also used the probe 2′7′-dichlorodihydrofluorescein diacetate (H2DCF-DA) to observe ROS accumulation in chloroplasts73. Briefly, PhANGs were transiently expressed in tobacco and harvested at 36 hpi. A solution of 10 μM H2DCF-DA in 10 mM MgCl2 buffer was injected into the infiltrated leaves using a syringe. For H2O2 detection in wheat protoplasts, TaHRP1, TaHRP1H/A, and TaPIR1 were transfected into wheat protoplasts via PEG-mediated transfection, following which the wheat protoplasts were harvested at 36 hpi and treated with 10 μM H2DCF-DA in 10 mM MgCl2 buffer. Chloroplast-derived ROS were visualized and imaged via confocal microscopy under laser excitation at 488 nm and emission at 530 nm.
Genomic DNA and RNA extraction and qRT-PCR analysis
Genomic DNA was extracted from the second leaves of two-leaf stage Fielder and transgenic plants using the CTAB method74. DNA from Pst-infected leaves was extracted at 5 dpi for fungal biomass analysis. Pst biomass was calculated using the cycle threshold (Ct) value of PstEF DNA against the Ct of wheat genomic elongation factor DNA (TaEF1). Total RNA was extracted from Pst-infected wheat leaves using MiniBEST Plant RNA Extraction Kit (Takara, Dalian, China) following the manufacturer’s instructions. First-strand cDNA synthesis was performed using the GoScript reverse transcription system (Promega Corp.). TaEF and PstEF were used to normalize the RNA levels of wheat leaves and Pst, respectively. Data from three independent biological replicates were analyzed using Student’s t-test.
Statistics and reproducibility
Image Lab (version 4.0) build 16 was used for DNA electrophoresis and WB data collection. The micrographs were collected by Leica LAS X Hardware Configurator Version 2020.6.0. MEME Suite 5.4.1 was used for analyzing conserved motifs. The sequence alignment was performed using http://multalin.toulouse.inra.fr/multalin/multalin.html. MEME Suite 5.4.1 was used for analyzing conserved motifs. All data analysis was performed using GraphPad 8.0 and is shown as means ± standard deviation (SD). The statistical analyses were performed using SPSS 26.0. The significant differences between experimental and control groups were determined by the two-tailed Student’s t-test, and one-way ANOVA. Analytical methods, n-values, and P values can be found in the figures or figure legends. All experiments in this study were performed two to and multiple times with similar results. No data were excluded from the analyses. The investigators were not blinded to allocation during experiments and outcome assessment.
Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.
Data availability
GenBank accession codes include XP_037425335 for TaPIR1 genes and XP_044353823 for TaHRP1 genes (https://www.ncbi.nlm.nih.gov/). DAP-seq data in this study has been uploaded to the NCBI Sequence Read Archive under the accessions PRJNA1066020. All data supporting the findings of this work are available in the paper, Supplementary Information files, and repository platform. Source data are provided with this paper.
Change history
27 February 2025
A Correction to this paper has been published: https://doi.org/10.1038/s41467-025-57387-4
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Acknowledgements
We thank Prof. Qixin Sun and Mingming Xin (China Agricultural University) for providing the pBUE411 vector used in this study. This study was supported by the National Natural Science Foundation of China (32102229) to Q.X., the Biotech Breeding Program of State Key Laboratory of Crop Gene Exploration and Utilization in Southwest China (SKL-ZY202233) to Q.X., the Sichuan Science and Program (2024NSFSC0405 and 2021YJ0298) to Q.X., and the Sichuan Science and Technology Program, China (2022ZDZX0014) to Y. Wei.
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Q.X., R.Z., and Y. Wei designed the research. R.Z., Y. Wu, and W.Y. conducted most of the experiments. X.Q. constructed vectors and performed the photosynthesis assay and BIFC assay. Q.W. and Y. Wu cultured tobacco and wheat plants. L.H., J.M., Y.Z., G.C., P.Q., Y.J., Q.J. and Y.Z. provided technical support. R.Z., X.W., Y. Wei, and Q.X. wrote and revised this manuscript.
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Zhang, R., Wu, Y., Qu, X. et al. The RING-finger ubiquitin E3 ligase TaPIR1 targets TaHRP1 for degradation to suppress chloroplast function. Nat Commun 15, 6905 (2024). https://doi.org/10.1038/s41467-024-51249-1
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DOI: https://doi.org/10.1038/s41467-024-51249-1
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