The ZmRACK1–ZmCDPK7–ZmAPX1 module regulates plant antiviral immunity

ABSTRACT

Calcium‐dependent protein kinases (CDPKs) in plants play crucial roles in mediating responses to both biotic and abiotic stresses. However, the molecular mechanism through which CDPKs regulate antiviral immunity in plants remains largely elusive. In this study, we identified ZmCDPK7 as a key player in response to various stimuli. Lines overexpressing ZmCDPK7 exhibited enhanced resistance to maize chlorotic mottle virus (MCMV) infection, while ZmCDPK7 knock‐out lines demonstrated increased sensitivity, indicating that ZmCDPK7 positively regulates maize immunity against MCMV. Furthermore, ZmCDPK7 interacts with ZmAPX1 and enhances its enzymatic activity by phosphorylating ZmAPX1 at Thr164, thereby reducing reactive oxygen species (ROS) production. The scaffold protein ZmRACK1 interacts with both ZmCDPK7 and ZmAPX1, facilitating the formation of a ternary complex that enhances ZmCDPK7‐mediated ZmAPX1 enzymatic activity. Lines overexpressing ZmAPX1 also showed increased resistance to MCMV infection, while ZmAPX1 mutant lines (Zmapx1 MU1 and Zmapx1 MU2) exhibited increased sensitivity. Additionally, CRISPR/Cas9‐mediated genome editing of ZmRACK1 in maize resulted in increased susceptibility to MCMV. To counteract plant defense, the MCMV protein P31 interacts with ZmCDPK7 and ZmRACK1, disrupting the interaction between ZmRACK1–ZmCDPK7 and ZmCDPK7–ZmAPX1, thereby blocking the ternary complex formation. Moreover, ZmCDPK7 is also implicated in defense against drought stress and corn stalk rot. Overall, our findings provide valuable insights into the molecular mechanisms by which the ZmRACK1–ZmCDPK7–ZmAPX1 module regulates plant immunity, identifying the potential new targets for the genetic control of maize viral diseases.

In maize, ZmRACK1 acts as a scaffold protein that bridges the cyclin‐dependent kinase ZmCDPK7 and the ascorbate peroxidase ZmAPX1, promoting ZmAPX1 activity to mitigate MCMV infection. Upon viral infection, P31 obstructs formation of the ZmRACK1‐ZmCDPK7‐ZmAPX1 complex, inhibiting ZmCDPK7‐mediated ZmAPX1 activity and promoting reactive oxygen species production to facilitate viral infection.

INTRODUCTION

Calcium is a ubiquitous intracellular second messenger in plant immune signaling, facilitating rapid responses that mediate plant responses to both biotic and abiotic stresses. In plants, calcium‐dependent protein kinases (CDPKs) are the sole calcium sensors that integrate both a sensor domain and a kinase domain, enabling them to decode Ca²⁺ signals into downstream signal transduction events (Yip Delormel and Boudsocq, 2019; Zhu et al., 2024b). CDPKs are implicated in nearly all aspects of the plant life cycle, from development to responses against biotic and abiotic stresses (Monaghan et al., 2014; Matschi et al., 2015; Hu et al., 2021; Su et al., 2024; Zhu et al., 2024a, 2024b). In maize, ZmCDPK39 negatively regulates quantitative resistance to gray leaf spot, northern leaf blight, and southern leaf blight (Zhu et al., 2024a). Additionally, two calcium‐dependent protein kinases, ZmCDPK35 and ZmCDPK37, enhance maize drought tolerance by activating ZmSLAC1 in guard cells (Li et al., 2022). However, the immune signaling pathway regulated by ZmCDPKs remains poorly understood.

Maize chlorotic mottle virus (MCMV) is a member of the genus Machlomovirus within the family Tombusviridae (Jiao et al., 2021). It can induce maize lethal necrosis (MLN) through synergistic infection with other viruses in the Potyviridae family, such as sugarcane mosaic virus (SCMV) (genus Potyvirus), leading to significant economic losses in various regions worldwide (Braidwood et al., 2018; Wu et al., 2024). MCMV possesses a positive‐sense, single‐stranded RNA genome that encodes seven proteins (Liu et al., 2022). P31, the pathogenicity determinant of MCMV, is crucial for successful viral infection (Jiao et al., 2021). Recent studies have shown that P31 hijacks maize catalase 1 to enhance viral multiplication and infection. Furthermore, P31 directly targets ZmPAO1 and counteracts the Zma‐miR167–ZmARF3/30 module‐mediated host defense (Liu et al., 2022). Current studies on the interaction between calcium‐dependent protein kinases (CDPKs) and plant viruses are limited. The active form of AtCDPK3 can restrict cell‐to‐cell propagation of potato virus X by phosphorylating the remorin StRem1.3 at residues S74, T86, and S91 (Perraki et al., 2018). The thiazole moiety TaTHI2 interacts with TaCDPK5 to suppress Chinese wheat mosaic virus (CWMV) infection by regulating ROS accumulation (Yang et al., 2023).

ROS play a crucial role in plant defense against viruses by integrating various signaling networks (Xu et al., 2024). The citrus tristeza virus (CTV) encoded p33 contributes to the induction of ROS accumulation and programmed cell death, thereby restricting CTV entry into phloem tissue and minimizing disease symptoms (Sun and Folimonova, 2019). Conversely, the wheat yellow mosaic virus (WYMV)‐encoded P1 inhibits ROS accumulation to facilitate viral infection. Overexpression of NbNAC1 prevents oxidative damage and reduces ROS production, enhancing plant resistance against viral pathogens (Zhu et al., 2025). Furthermore, while ROS promotes certain viral infections and transmission, a previous report found that lower ROS accumulation inhibited MCMV infection (Liu et al., 2022). The barley yellow dwarf virus (BYDV) movement protein interacts with both wheat catalases (CATs) and the 26S proteasome ubiquitin receptor PSMD2 to facilitate the 26S proteasome‐mediated degradation of CATs. This degradation promotes bursts of reactive oxygen species, leading to enhanced BYDV accumulation and the development of disease symptoms (Tian et al., 2024). Additionally, the replication of red clover necrotic mosaic virus (RCNMV), a positive‐sense RNA virus, requires NbRBOHB‐mediated ROS production (Hyodo et al., 2019).

To investigate the role of ZmCDPKs in plant antiviral immunity against MCMV infection, we identified ZmCDPK7, a calcium‐dependent protein kinase that responds to various biotic and abiotic stresses. Here, we report that ZmCDPK7 interacts with both ZmRACK1 and ZmAPX1, phosphorylating ascorbate peroxidase and enhancing ZmAPX1 activity to regulate ROS production. Knockout mutants of ZmCDPK7 and ZmRACK1, as well as ZmAPX1 mutant lines, exhibit increased susceptibility to MCMV infection. Furthermore, MCMV P31 disrupts the interaction between ZmCDPK7–ZmRACK1 and ZmCDPK7–ZmAPX1, thereby inhibiting the ZmRACK1–ZmCDPK7–ZmAPX1‐mediated defense and promoting viral infection. Our study elucidates the molecular mechanism by which the ZmRACK1–ZmCDPK7–ZmAPX1 regulatory module mediates immunity against MCMV.

RESULTS

Identification of ZmCDPK7 in response to various stimuli

CDPKs are involved in immune signaling upon the perception of external stimuli (Matschi et al., 2015; Hu et al., 2021; Mou et al., 2024). We analyzed the evolutionary relationships of the ZmCDPK gene family. To accurately delineate the phylogenetic relationships, we obtained CDPK gene sequences from rice and wheat in GenBank. The results revealed a total of 48 members of the ZmCDPK gene family, categorized into four subfamilies (Figure S1A). We analyzed public maize transcriptome data, elucidating expression profiles during infection by major maize pathogens. The findings indicated that most family members exhibited expression changes induced by varying degrees of regulatory influences from different biotic stresses (Figure S1B). Notably, ZmCDPK7 exhibited significantly increased expression levels in response to infections by Rhopalosiphum maidis, Ustilago maydis, and Fusarium graminearum (Figure S1C). Importantly, CDPK5, which is a counterpart of ZmCDPK7, exhibited significant upregulation in both wheat and rice, specifically under Fusarium graminearum infection in wheat and Magnaporthe oryzae infection in rice. This suggests a conserved role in stress that extends beyond maize (Figure S1D). Based on these findings, we selected ZmCDPK7 for further study. Given the ambiguous role of ZmCDPKs in viral infection, we examined the expression of ZmCDPK7 in the transcriptome of mock‐treated and MCMV‐infected maize leaves. The results indicated that ZmCDPK7 was transcriptionally up‐regulated following MCMV infection (Figure S1E). Furthermore, we assessed the expression of ZmCDPK7 in response to MCMV and SCMV at different time points. The findings demonstrated that the expression of ZmCDPK7 was induced during the infections of both SCMV and MCMV (Figure S1F–G).

ZmCDPK7 positively regulates resistance to MCMV

To elucidate the function of ZmCDPK7 during MCMV infection, we generated knockout mutants of the endogenous ZmCDPK7 in the B104 background using CRISPR‐Cas9‐mediated genome editing. We identified two independent T1 knockout lines, Zmcdpk7 KO1 and Zmcdpk7 KO2, which contain 31‐ and 32‐nt deletions, respectively (Figure 1A). Both Zmcdpk7 KO1 and Zmcdpk7 KO2 lines were inoculated with MCMV. At 7 d post‐inoculation (dpi), the Zmcdpk7 KO1 and Zmcdpk7 KO2 lines exhibited more severe chlorosis and mottling symptoms compared to control plants (Figure 1B). We conducted RT‐qPCR and western blotting assays, and the results indicated that MCMV accumulation was increased in Zmcdpk7 KO1 and Zmcdpk7 KO2 plants compared to control plants (Figure 1C, D). To further confirm the function of ZmCDPK7 during MCMV infection, we employed a CMV‐based virus‐induced gene silencing (VIGS) system to silence ZmCDPK7. At 10 dpi, maize plants were inoculated with MCMV. After 7 d, we assessed the silencing efficiency of ZmCDPK7 using RT‐qPCR and observed significant downregulation of ZmCDPK7 expression in CMV‐ZmCDPK7‐infected maize plants (Figure S2A). The ZmCDPK7‐silenced plants exhibited more pronounced chlorosis and mottling symptoms compared to the non‐silenced control plants following MCMV infection (Figure S2B). Subsequently, we next analyzed the viral accumulation of MCMV using both RT‐qPCR and western blotting. The results indicated that MCMV accumulation was higher in ZmCDPK7‐silenced plants compared to the control plants (Figure S2C, D).

Figure 1.

ZmCDPK7 positively regulates MCMV resistance in maize

(A) Diagram of the mutations in the ZmCDPK7 mutants. The blue and yellow rectangles and black line indicate the untranslated regions (UTRs), exons, and introns of ZmCDPK7, respectively. The positions of sgRNA1 and sgRNA2 are indicated by black arrows. The sequences of sgRNA1 and sgRNA2 are shown below the diagram. (B) Phenotypes of B104, Zmcdpk7 KO1, and Zmcdpk7 KO2 plants infected with MCMV at 7 dpi. Scale bar = 1 cm. (C) MCMV coat protein (CP) transcript levels in B104, Zmcdpk7 KO1, and Zmcdpk7 KO2 plants, quantified by RT‐qPCR. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was assessed using Student's t‐test (*P < 0.05). (D, H) MCMV CP accumulation in systemically infected leaves analyzed by a western blot using an MCMV CP‐specific antibody. (E) ZmCDPK7 expression levels in B104, ZmCDPK7 OE#1, and ZmCDPK7 OE#2 plants determined by quantitative RT‐PCR. ZmUBI was used as an internal reference gene. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was evaluated using Student's t‐test (**P < 0.01). (F) Phenotypes of B104, ZmCDPK7 OE#1, and ZmCDPK7 OE#2 plants infected with MCMV at 7 dpi. Scale bar = 1 cm. (G) MCMV CP transcript levels in B104, ZmCDPK7 OE#1, and ZmCDPK7 OE#2 plants measured by RT‐qPCR. The amplification of ZmUBI served as an internal control. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was determined by Student's t‐test (*P < 0.05).

To further investigate the role of ZmCDPK7, transgenic maize in the B104 background was generated that overexpresses a GFP‐tagged ZmCDPK7 under the ubiquitin promoter. The expression of ZmCDPK7 was upregulated in the ZmCDPK7 OE#1 and ZmCDPK7 OE#2 T2 transgenic lines (Figure 1E). The ZmCDPK7 OE#1, ZmCDPK7 OE#2, and B104 were inoculated with MCMV. At 7 dpi, the ZmCDPK7 OE#1 and ZmCDPK7 OE#2 lines exhibited milder symptoms compared to the control plants (Figure 1F). Additionally, RT‐qPCR and western blot results demonstrated lower accumulation levels of MCMV CP RNA and protein in the ZmCDPK7 OE#1 and ZmCDPK7 OE#2 lines inoculated with MCMV (Figure 1G, H).

ZmCDPK7 enhances the enzymatic activity of ZmAPX1 by phosphorylating it at Thr164

To further identify proteins regulated by ZmCDPK7, a yeast two‐hybrid screening was conducted, and ZmAPX1 was identified as a candidate interacting protein. The interaction between ZmCDPK7 and ZmAPX1 was confirmed using the yeast two‐hybrid (Y2H) system (Figure 2A). To validate the interaction in vivo, we performed a bimolecular fluorescence complementation (BiFC) assay (Figure 2B). YFP fluorescence was observed in the combinations of ZmCDPK7 and ZmAPX1, with no fluorescence detected in negative controls. Additionally, a co‐immunoprecipitation assay (Co‐IP) was performed, in which ZmAPX1 was fused with GFP and ZmCDPK7 with Myc for co‐expression in N. benthamiana tissues. Co‐IP results demonstrated that ZmAPX1‐GFP co‐precipitated with ZmCDPK7‐Myc, but not with the GUS‐Myc control (Figure 2C). CDPKs have been reported to be involved in ROS synthesis (Yu et al., 2024a). To further investigate the role of ZmCDPK7 in regulating ROS and viral infection, we analyzed ROS levels in Zmcdpk7 KO1 and Zmcdpk7 KO2 lines using DAB staining and assessed H₂O₂ accumulation, which was found to be elevated in both Zmcdpk7 KO1 and Zmcdpk7 KO2 lines (Figure S3A, B).

Figure 2.

ZmCDPK7 interacts with ZmAPX1 and enhances its activity via phosphorylation at Ser164

(A) Y2H assays showing the interaction between ZmCDPK7 and ZmAPX1. ADT + BD53a and ADT + BDLam served as positive and negative controls, respectively. (B) BiFC assay showing the interaction between ZmCDPK7 and ZmAPX1 in N. benthamiana leaves. Scale bar = 50 μm. (C) Co‐IP assay to assess the ZmCDPK7–ZmAPX1 interaction. Nicotiana benthamiana leaves were co‐infiltrated with Agrobacterium tumefaciens cells harboring expression vectors for ZmCDPK7‐Myc and ZmAPX1‐GFP, or GUS‐Myc and ZmAPX1‐GFP. (D) In vitro phosphorylation assay. The phosphorylation reactions were conducted using ZmCDPK7‐His as the kinase and ZmAPX1‐His as the substrate, followed by immunoblotting with an anti‐phospho‐serine/threonine (anti‐Phospho) antibody. (E) In planta phosphorylation assay. Nicotiana benthamiana leaves were co‐infiltrated with A. tumefaciens cells harboring expression vectors for ZmCDPK7‐Myc and ZmAPX1‐GFP, or GUS‐Myc and ZmAPX1‐GFP. (F) In planta phosphorylation assay. Nicotiana benthamiana leaves were co‐infiltrated with Agrobacterium tumefaciens cells harboring expression vectors for ZmCDPK7^ΔKD‐Myc and ZmAPX1‐GFP, or ZmCDPK7‐Myc and ZmAPX1‐GFP. (G) APX enzyme activity assays. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was assessed using Student's t‐test (**P < 0.01). (H) Measurement of H₂O₂ content. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was assessed using Student's t‐test (*P < 0.05). (I) The phosphorylation site of ZmAPX1. Site‐directed mutagenesis was used to generate three variants: ZmAPX1^T59A, ZmAPX1^T164A, and ZmAPX1^T59A/T164A. (J) APX enzyme activity assays. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was assessed using Student's t‐test (*P < 0.05). (K) Measurement of H₂O₂ content. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was assessed using Student's t‐test (**P < 0.01).

Given that ZmCDPK7 interacts with ZmAPX1, we propose that ZmCDPK7 likely phosphorylates ZmAPX1 through this interaction. To validate this hypothesis, in vitro kinase activity assays were conducted. A ZmCDPK7‐His fusion protein was shown to phosphorylate a ZmAPX1‐His substrate but not with MBP‐His in vitro, evidenced by immunoblotting with an anti‐phospho‐serine/threonine (anti‐pSer/pThr) antibody (Figure 2D). When a GFP‐fused ZmAPX1 construct (ZmAPX1‐GFP) and a Myc‐fused ZmCDPK7 construct (ZmCDPK7‐Myc) were co‐expressed in N. benthamiana leaves, the phosphorylation level of ZmAPX1 was significantly increased, indicating that ZmAPX1 is phosphorylated by ZmCDPK7 in vivo (Figure 2E). Moreover, we observed that the phosphorylation level of ZmAPX1 was reduced when coexpressing with the kinase domain deletion mutant of ZmCDPK7 (ZmCDPK7^ΔKD) (Hrabak et al., 1996; Figure 2F). The Lys163 residue has been identified as a key phosphorylation site (Zhou et al., 2007), prompting us to perform site‐directed mutagenesis to replace it with Arg in ZmCDPK7. Subsequent analysis revealed that the ZmCDPK7^K163R mutant failed to phosphorylate ZmAPX1 in vitro (Figure S4). To further verify the regulatory effect of ZmCDPK7 on ZmAPX1, we assessed APX enzymatic activity in vivo. We observed that APX enzymatic activity was higher in the presence of ZmCDPK7 (Figure 2G), while ZmCDPK7^ΔKD showed no obvious impact on the APX enzymatic activity, indicating that the phosphorylation of ZmAPX1 by ZmCDPK7 is essential for its enzymatic activity. Moreover, we evaluated the accumulation of H₂O₂. The expression of ZmCDPK7 inhibited the accumulation of H₂O₂ compared to the GUS control, while ZmCDPK7^ΔKD exhibited no difference (Figure 2H).

To identify the residues that are phosphorylated by ZmCDPK7 in ZmAPX1, two Thr residues were found via online prediction: Thr59 and Thr164. When ZmAPX1^T164A‐GFP, ZmAPX1^T59A/T164A‐GFP, and ZmCDPK7‐Myc were co‐expressed in N. benthamiana leaves, the phosphorylation level of ZmAPX1 was significantly decreased, whereas ZmAPX1^T59A‐GFP expression had no obvious effect on the phosphorylation level of ZmAPX1, indicating that Thr164 is likely the phosphorylation site in ZmAPX1 by ZmCDPK7 (Figure 2I). We further tested the effect of the phosphorylation site mutation on APX enzymatic activity in vivo. We observed that ZmAPX1^T164A and ZmAPX1^T59A/T164A reduced their enzymatic activity (Figure 2J) and hence promoted H₂O₂ accumulation (Figure 2K). The results further confirmed that ZmAPX1 phosphorylation by ZmCDPK7 is indispensable for the enzymatic activity.

ZmAPX1 contributes to the defense against MCMV

To investigate the role of ZmAPX1 in viral infection, the expression of ZmAPX1 during MCMV infection was assessed using RT‐qPCR (Figure S5A). ZmAPX1 was found to be upregulated during MCMV infection in maize. DAB staining indicated low accumulation of ROS in MCMV‐infected leaves at both 2 dpi and 10 dpi (Figure S5B). A CMV‐based VIGS system was employed to elucidate the function of ZmAPX1. At 10 dpi, maize plants were inoculated with MCMV, and after 7 d, the silencing efficiency of ZmAPX1 was confirmed via RT‐qPCR, revealing significant downregulation of ZmAPX1 expression in CMV‐ZmAPX1‐infected maize plants (Figure S6A). ZmAPX1‐silenced plants exhibited increased susceptibility to MCMV compared to the non‐silenced control (Figure S6B), with increased viral accumulation in CMV‐ZmAPX1‐silenced plants (Figure S6C, D).

Further, we utilized a ZmAPX1 mutant with mutator transposon insertions in the 5′ UTR. At 7 dpi, the ZmAPX1 mutant lines (zmapx1 MU1 and zmapx1 MU2) exhibited more obvious symptoms compared to control plants (Figure 3A). MCMV accumulation was significantly higher in zmapx1 MU1 and zmapx1 MU2 lines than in the control plants, as evidenced by both RT‐qPCR and western blotting analysis (Figure 3B, C). To further elucidate the role of ZmAPX1, ZmAPX1‐overexpressing lines (ZmAPX1 OE#1 and ZmAPX1 OE#2) were inoculated with MCMV. At 7 dpi, ZmAPX1 OE#1 and ZmAPX1 OE#2 lines exhibited milder symptoms and showed resistance to MCMV infection (Figure 3D). The RT‐qPCR and western blot analyses indicated lower accumulation levels of MCMV CP RNA and protein in ZmAPX1 OE#1 and ZmAPX1 OE#2 lines inoculated with MCMV (Figure 3E, F). DAB staining further demonstrated that overexpression of ZmAPX1 inhibited ROS accumulation, while knockout of ZmAPX1 promoted ROS accumulation during MCMV infection (Figure S7A, B).

Figure 3.

ZmAPX1 contributes to the defense against MCMV

(A) Phenotypes of WT, zmapx1 MU1, and zmapx1 MU2 plants infected with MCMV at 7 dpi. Scale bar = 1 cm. (B) The levels of MCMV CP transcripts, quantified by RT‐qPCR in zmapx1 MU1 and zmapx1 MU2 lines, and WT. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was determined by Student's t‐test (**P < 0.01). (C) The accumulation of MCMV CP in systemically infected leaves of WT, zmapx1 MU1, and zmapx1 MU2 plants. (D) Phenotypes of WT, ZmAPX1 OE#1, and ZmAPX1 OE#2 plants infected with MCMV at 7 dpi. Scale bar = 1 cm. (E) MCMV CP transcript levels in WT, ZmAPX1 OE#1, and ZmAPX1 OE#2 plants measured by RT‐qPCR. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was determined by Student's t‐test (**P < 0.01). (F) The accumulation of MCMV CP in systemically infected leaves of WT, ZmAPX1 OE#1, and ZmAPX1 OE#2 plants.

ZmRACK1 interacts with ZmCDPK7 and ZmAPX1 and promotes ZmCDPK7‐regulated ZmAPX1 enzymatic activity via enhancing the interaction

ZmRACK1 was identified as another candidate interacting protein of ZmCDPK7 through yeast two‐hybrid screening (Table S2). Receptor for activated C kinase 1 (RACK1) is a highly conserved scaffold protein involved in various signaling pathways (Nakashima et al., 2008). In this study, we investigated the potential direct binding of ZmRACK1 to ZmCDPK7 and ZmAPX1. Y2H assays demonstrated that ZmRACK1 interacts with both ZmCDPK7 and ZmAPX1 (Figure 4A, D). These interactions were subsequently verified by BiFC assays conducted in N. benthamiana leaves (Figure 4B, E). Furthermore, the interactions between ZmRACK1 and ZmCDPK7, as well as ZmRACK1 and ZmAPX1, were validated using Co‐IP assays (Figure 4C, F). To investigate whether ZmRACK1, ZmCDPK7, and ZmAPX1 may interact with one another in planta, we performed a Co‐IP assay in which ZmRACK1 was fused to GFP, ZmCDPK7 to Myc, and ZmAPX1 to Flag for co‐expression in N. benthamiana tissues. The results indicated that ZmRACK1, ZmCDPK7, and ZmAPX1 likely interact with each other, forming a protein complex within N. benthamiana leaf cells (Figure 4G).

Figure 4.

ZmRACK1 interacts with ZmCDPK7 and ZmAPX1 and enhances their interaction

(A, D) Y2H assays showing the interaction between ZmCDPK7–ZmRACK1 and ZmAPX1–ZmRACK1. The combinations of ADT + BD53a and ADT + BDLam served as positive and negative controls, respectively. (B, E) BiFC assay showing the interaction between ZmCDPK7–ZmRACK1 and ZmAPX1–ZmRACK1 in Nicotiana benthamiana leaves. Scale bar = 50 μm. (C, F) Co‐IP assay to assess the ZmCDPK7–ZmRACK1 and ZmAPX1–ZmRACK1 interaction in vivo. (G) Co‐IP assay to assess the interaction among ZmCDPK7, ZmRACK1, and ZmAPX1 in vivo. (H, I) BiFC assay to assess the interaction between ZmCDPK7 and ZmAPX1, with or without ZmRACK1. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 10) was assessed using Student's t‐test (**P < 0.01). Scale bar = 50 μm. (J) Co‐IP assay to assess the effect of ZmRACK1 on the interaction between ZmCDPK7–ZmAPX1 in vivo. (K) In planta phosphorylation assay. Nicotiana benthamiana leaves were co‐infiltrated with Agrobacterium tumefaciens cells harboring expression vectors for ZmCDPK7‐Myc, ZmAPX1‐GFP, ZmRACK1‐Flag, or GUS‐Flag. Two days later, phosphorylated ZmAPX1 proteins were immunoprecipitated using anti‐GFP agarose beads (IP: anti‐GFP) and immunoblotted with anti‐Phospho antibodies. Samples before immunoprecipitation were analyzed by immunoblotting using anti‐GFP, anti‐Myc, or anti‐Flag antibodies.

To assess the role of ZmRACK1 in regulating the enzymatic activity of ZmAPX1 via ZmCDPK7, we co‐expressed ZmRACK1 with ZmCDPK7 and found that ZmRACK1 expression enhanced ZmAPX1 enzymatic activity regulated by ZmCDPK7 while reducing H₂O₂ accumulation (Figure S8A, B). Given that ZmRACK1 positively influences ZmCDPK7 function, we sought to determine whether ZmRACK1 could modulate the interaction between ZmCDPK7 and ZmAPX1. BiFC was conducted to examine the interaction between ZmCDPK7 and ZmAPX1 in the presence of ZmRACK1, revealing increased YFP fluorescence in the combinations of ZmCDPK7‐nYFP and ZmAPX1‐cYFP with ZmRACK1 expression (Figure 4H, I). Additionally, a Co‐IP assay assessing the interaction between ZmCDPK7 and ZmAPX1 at varying concentrations of ZmRACK1 showed that higher ZmRACK1 concentration corresponded with an increased amount of ZmCDPK7 bound by ZmAPX1‐GFP. In contrast, increasing GUS concentration did not affect the interaction between ZmCDPK7 and ZmAPX1 (Figure 4J). Furthermore, to investigate the influence of ZmRACK1 expression on the phosphorylation of ZmAPX1 mediated by ZmCDPK7 in N. benthamiana, we expressed ZmRACK1 together with ZmAPX1 and ZmCDPK7. Our findings indicate that the transient expression of ZmRACK1 enhanced ZmCDPK7‐mediated phosphorylation of ZmAPX1 (Figure 4K).

ZmRACK1 positively regulates resistance to MCMV

To further elucidate the role of ZmRACK1, we generated knockout mutants of the endogenous ZmRACK1 (zmrack1 KO1 and zmrack1 KO2) in the B104 background through CRISPR‐Cas9‐mediated genome editing (Figure 5A). The zmrack1 KO1, zmrack1 KO2, and B104 were inoculated with MCMV. At 7 dpi, the Zmrack1 KO1 and Zmrack1 KO2 lines exhibited increased susceptibility to MCMV compared to the control plants (Figure 5B). The accumulation of MCMV was significantly increased in zmrack1 KO1 and zmrack1 KO2 plants compared to control plants, as demonstrated by both RT‐qPCR and western blotting analyses (Figure 5C, D). Additionally, the levels of H₂O₂ were markedly elevated in zmrack1 KO1 and zmrack1 KO2 lines relative to wild‐type lines (Figure S9A, B). Moreover, a CMV‐based VIGS system was employed. At 10 dpi, maize plants were further inoculated with MCMV. After 7 d, we assessed the silencing efficiency of ZmRACK1 using RT‐qPCR and observed significant downregulation of ZmRACK1 expression in CMV‐ZmRACK1‐infected maize plants (Figure S10A). Notably, ZmRACK1‐silenced plants exhibited increased susceptibility to MCMV compared to the non‐silenced control. Viral accumulation was increased in CMV‐ZmRACK1‐silenced plants, as confirmed by both RT‐qPCR and western blotting analyses (Figure S10B–D).

Figure 5.

ZmRACK1 positively regulates maize resistance to MCMV

(A) Diagram of the mutations in the zmrack1 mutants. The blue and yellow rectangles and black line indicate the untranslated regions (UTRs), exons, and introns of ZmRACK1, respectively. The positions of sgRNA1 and sgRNA2 are indicated by black arrows. The sequences of sgRNA1 and sgRNA2 are shown below the diagram. Two independent knockout lines, zmrack1 KO1 (with a 164 bp deletion in zmrack) and zmrack1 KO2 (with a 2 bp deletion and 1 bp insertion in zmrack1), were generated by CRISPR‐Cas9 genome editing in the B104 inbred line background. (B) Phenotypes of B104, zmrack1 KO1, and zmrack1 KO2 plants infected with MCMV at 7 dpi. Scale bar = 1 cm. (C) MCMV CP transcript levels in B104, zmrack1 KO1, and zmrack1 KO2 plants were measured by RT‐qPCR. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was determined using Student's t‐test (**P < 0.01). (D) MCMV CP accumulation in systemically infected leaves determined by western blotting using an MCMV CP‐specific antibody. (E) ZmRACK1 expression in ZmRACK1‐silenced plants and control plants assessed by RT‐qPCR. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was determined by Student's t‐test (**P < 0.01). (F) Phenotypes of B104, ZmCDPK7 OE#1, and ZmCDPK7 OE#2 plants with or without ZmRACK1 silencing infected with MCMV at 7 dpi. Scale bar = 1 cm. (G, H) MCMV CP transcripts and CP levels measured in B104, ZmCDPK7 OE#1, and ZmCDPK7 OE#2 plants with or without ZmRACK1 silencing, measured by RT‐qPCR and western blot, respectively. Amplification of ZmUBI served as the internal control. Bars represent the means ± SE. Statistical significance between the treatments (n ≥ 5) was determined by Student's t‐test (*P < 0.05).

Furthermore, we silenced ZmRACK1 in ZmCDPK7 OE#1 and ZmCDPK7 OE#2 lines (Figure 5E). The results indicated that transient silencing of ZmRACK1 did not significantly affect the antiviral function conferred by ZmCDPK7 overexpression, suggesting that ZmRACK1 likely functions upstream of ZmCDPK7 (Figure 5F–H).

P31 interacts with ZmCDPK7 and ZmRACK1 and interferes with the interaction of the complex ZmRACK1–ZmCDPK7–ZmAPX1

During the arms race between virus and host, viruses encode proteins to counteract plant immunity (Chen et al., 2023). A Y2H assay was performed to identify viral proteins that interact with ZmCDPK7. The results indicated that P31 interacted with ZmCDPK7 in yeast (Figures 6A, S11). These interactions were subsequently verified through BiFC assays in N. benthamiana leaves. YFP fluorescence was observed in the combination of ZmCDPK7‐cYFP and P31‐nYFP, but not in the control combinations (Figure 6B). The interaction was further validated using a Co‐IP assay, which demonstrated that ZmCDPK7‐GFP co‐precipitated with P31‐Myc, but not with the GUS‐Myc control (Figure 6C). Additionally, we confirmed the interaction between ZmRACK1 and P31 through Y2H (Figure 6D). This interaction was also verified by BiFC assays (Figure 6E). The Co‐IP assay results demonstrated that P31‐Myc co‐precipitated with ZmRACK1‐GFP, but not with the GFP control, in Nicotiana benthamiana (Figure 6F). The interaction between P31 and ZmAPX1 was unfound by BiFC assays in N. benthamiana leaves (Figure S12). To evaluate the role of P31 in ZmCDPK7‐mediated immunity, a BiFC assay was conducted to investigate the interaction between ZmCDPK7 and ZmAPX1 in the presence of P31. The YFP fluorescence was diminished in the combinations of ZmCDPK7 and ZmAPX1 with the expression of P31 (Figure 6G, H). Furthermore, a Co‐IP assay of ZmCDPK7 and ZmAPX1 was performed using varying concentrations of P31. We observed that increasing the concentration of P31 gradually reduced the amount of ZmAPX1‐Flag bound to ZmCDPK7‐Myc, whereas the increase of the GFP control did not affect the interaction between ZmCDPK7 and ZmAPX1 (Figure 6I). Additionally, in the presence of P31, YFP fluorescence was also reduced in the combinations of ZmCDPK7‐nYFP and ZmRACK1‐cYFP (Figure 6J–K). A Co‐IP assay of ZmCDPK7 and ZmRACK1 with different concentrations of P31 revealed that higher P31 concentrations led to a gradual decrease in the amount of ZmRACK1 bound by ZmCDPK7‐Myc. The increase in GFP control did not influence the interaction between ZmCDPK7 and ZmRACK1 (Figure 6L). Moreover, we found that P31 expression inhibited the phosphorylation of ZmAPX1 (Figure 6M).

Figure 6.

P31 interacts with ZmCDPK7 and interferes with its interaction with ZmAPX1

(A) Y2H assay showing interactions between ZmCDPK7 and MCMV‐encoded proteins. (B, E) BiFC assay showing the interactions between ZmCDPK7‐P31 and ZmRACK1‐P31 in Nicotiana benthamiana. Scale bar = 50 μm. (C, F) Co‐IP assay to assess the interaction of ZmCDPK7‐P31 and ZmRACK1‐P31 in vivo. (D) Y2H assay showing the interactions between P31 and ZmRACK1. ADT + BD53a and ADT + BDLam served as positive and negative controls, respectively. (G, H) BiFC assay showing the interaction between ZmCDPK7 and ZmAPX1, with and without P31. Scale bar = 50 μm. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 10) was determined by Student's t‐test (**P < 0.01). (I) Co‐IP assay to assess the ZmCDPK7‐ZmAPX1 interaction, with and without P31. (J, K) BiFC assay showing the interaction between ZmCDPK7 and ZmRACK1, with and without P31. Scale bar = 50 μm. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 10) was assessed using Student's t‐test (**P < 0.01). (L) Co‐IP assay to assess the ZmCDPK7–ZmRACK1 interaction, with and without P31. (M) In planta phosphorylation assay. Nicotiana benthamiana leaves were co‐infiltrated with Agrobacterium tumefaciens cells harboring expression vectors for ZmCDPK7‐Myc and ZmAPX1‐GFP alongside P31‐Myc or GUS‐Myc. Two days later, phosphorylated ZmAPX1 proteins were immunoprecipitated using anti‐GFP agarose beads (IP: anti‐GFP) and immunoblotted with anti‐Phospho antibodies. Samples before immunoprecipitation were analyzed by immunoblotting using anti‐GFP, anti‐Myc, or anti‐Flag antibodies.

ZmCDPK7 participates in plant response to abiotic and biotic stresses

Previous studies have reported that oilseed rape CDPK5 positively regulates drought tolerance (Cheng et al., 2022). To investigate the role of ZmCDPK7 under drought stress, two independent transgenic knockout lines, Zmcdpk7 KO1 and Zmcdpk7 KO2, were selected for stress tolerance assays. Under drought conditions, the Zmcdpk7 KO1 and Zmcdpk7 KO2 lines exhibited evident wilting and leaf rolling, while wild‐type plants showed only moderate wilting (Figure 7A, B). Furthermore, both Zmcdpk7 KO1, Zmcdpk7 KO2, and B104 exhibited a significant increase in MDA (malondialdehyde) content and electrolyte leakage, with levels higher in Zmcdpk7 KO1 and Zmcdpk7 KO2 than in B104 (Figure 7C, D). Additionally, H₂O₂ content was elevated in Zmcdpk7 KO1 and Zmcdpk7 KO2 compared to B104 (Figure 7E). These findings support the observed greater susceptibility of Zmcdpk7 knockout mutants to drought stress. Furthermore, we assessed the resistance of Zmcdpk7 KO1 and Zmcdpk7 KO2 lines to F. verticillioides (Fv), a major pathogen causing maize stalk rot. The observed symptom demonstrated that the two Zmcdpk7 knockout lines, KO1 and KO2, exhibited increased susceptibility to F. verticillioides (Figure 7F, G).

Figure 7.

ZmCDPK7 participates in plant responses to abiotic and biotic stresses

(A, B) Phenotypes of B104, Zmcdpk7 KO1, and Zmcdpk7 KO2B104 plants after 4 d of drought stress. Scale bar = 1 cm. (C–E) Electrolyte leakage (C), MDA contents (D), and H₂O₂ contents (E) in the leaves of B104, Zmcdpk7 KO1, and Zmcdpk7 KO2 plants. Scale bar = 1 cm. Bars represent the means ± SE. Statistical significance between the two treatments (n ≥ 5) was determined by Student's t‐test (*P < 0.05; **P < 0.01). (F, G) Symptoms in roots and stems of B104, Zmcdpk7 KO1, and Zmcdpk7 KO2 plants following inoculation with Fv. Scale bar = 1 cm.

DISCUSSION

As calcium sensors, CDPKs play crucial roles in regulating plant responses to environmental stimuli such as drought, hypoxia, salinity, and heat stress, as well as pathogen attack (Hyodo et al., 2017; Zhao et al., 2021; Sun et al., 2022; Mou et al., 2024; Su et al., 2024; Yu et al., 2024b; Zhu et al., 2024b). However, the role of CDPKs in plant viral infection remains poorly understood. In this study, we demonstrated that ZmCDPK7 regulates plant defense against MCMV by phosphorylating ascorbate peroxidase and enhancing ZmAPX1 activity. Additionally, ZmRACK1 promotes ZmCDPK7‐mediated ZmAPX1 activity through direct interaction with ZmCDPK5. During viral infection, the MCMV protein P31 interferes with the interactions between ZmCDPK7 and ZmRACK1, as well as between ZmCDPK7 and ZmAPX1, thereby inhibiting the ZmCDPK7–ZmRACK1–ZmAPX1‐mediated defense mechanism. Consequently, our findings elucidate a maize ZmCDPK7–ZmRACK1–ZmAPX1 module that mediates resistance to viral infection. Furthermore, we have identified a novel role for the viral protein P31 in the immune response pathway.

Among the 48 ZmCDPKs identified in maize, ZmCDPK39 has been reported as the first example of a ZmCDPK involved in a biotic stress response (Zhu et al., 2024b). ZmCDPK39 negatively regulates plant immunity. CDPKs from other plant species, such as AtCDPK28 and OsCDPK4, have been reported to negatively regulate plant immunity (Monaghan et al., 2014; Bundó and Coca, 2016; Wang et al., 2018). In this study, we demonstrated that ZmCDPK7 positively regulates immune responses against viral diseases in maize. ZmCDPK7 is transcriptionally induced upon viral infection. ZmCDPK7 knockout mutants and transiently silenced plants exhibited increased susceptibility to MCMV compared to wild‐type plants. Similar to ZmCDPK7, OsCDPK17 positively regulates immune responses and resistance against bacterial diseases in rice (Mou et al., 2024). Additionally, AtCDPK1 overexpression confers broad‐spectrum resistance against fungal and bacterial pathogens (Singh et al., 2017).

RACK1 functions as a scaffold protein and interacts with various proteins involved in biological processes, including development, miRNA biogenesis, hormone signaling, protein translation, and stress responses (Islas‐Flores et al., 2015; Hyodo et al., 2019). NbRACK1 has been shown to be recruited by the p27 viral replication protein into aggregated structures for the replication of red clover necrotic mosaic virus (RCNMV) (Hyodo et al., 2019) via ROS production. OsRACK1A interacts with key regulators of plant immunity and contributes to ROS bursts and resistance against the rice blast fungus (Nakashima et al., 2008; Yu et al., 2024b). In this study, we found that ZmRACK1 acts as a scaffold protein and functions in maize immunity by forming an immune complex with ZmCDPK7and ZmAPX1 to enhance the ZmAPX1 activity. The increase in ZmAPX1 activity leads to reduced low H₂O₂ accumulation and enhanced resistance against MCMV. A previous study found that ZmAPX1 overexpression increased resistance against Bipolaris maydis (Zhang et al., 2022).

ROS play crucial roles in viral infection and function as a double‐edged sword (Xu et al., 2024). The accumulation of ROS influences the redox state of numerous proteins and aids plants in developing systemic resistance against plant viruses (Sun and Folimonova, 2019; Zhu et al., 2025). In our study, however, ROS production facilitates MCMV infection. Liu et al. (2022) previously found that reduced ROS accumulation inhibited MCMV infection, which is consistent with our findings. Additionally, ROS generation enhances BYDV accumulation and the development of disease symptoms (Tian et al., 2024). Further studies are needed to clarify the role of ROS in promoting viral infection.

In summary, we have elucidated the molecular mechanism by which ZmCDPK7 positively regulates plant immunity through the ZmRACK1–ZmCDPK7–ZmAPX1 functional module (Figure S13). ZmRACK1 acts as a scaffold protein that connects ZmCDPK7 and ZmAPX1, enhancing ZmCDPK7‐mediated ZmAPX1 activity to mitigate viral infection. Upon viral infection, P31 disrupts the interaction between ZmCDPK7–ZmRACK1 and ZmCDPK7–ZmAPX1, thereby inhibiting ZmCDPK7‐mediated ZmAPX1 activity and promoting viral infection.

MATERIALS AND METHODS

Plant materials

Nicotiana benthamiana plants and maize (Zea mays L.) inbred lines B73 and B104 were cultivated in growth chambers under controlled conditions (24°C day/22°C night, 16 h photoperiod). All ZmCDPK7 and ZmRACK1 transgenic plants and mutants were generated by Weimi Biotechnology Company Ltd. The constructs were transformed into the inbred line B104, and T1 plants with stable expression or knockout were used in this study. The ZmAPX1 transgenic plants and mutants were kindly provided by Professor Liuji Wu from Henan Agricultural University.

Virus inoculation

The MCMV inoculum was derived from the full‐length cDNA clone pMCM41, which was kindly provided by Dr. Kay Scheets from Oklahoma State University. The SCMV‐BJ isolate was obtained from established sources (Fan et al., 2003). For both viruses, crude sap was prepared by homogenizing infected leaf tissue in 0.01 M phosphate buffer (49:51 v/v 0.01 M KH₂PO₄:Na₂HPO₄, pH 7.0) at a ratio of 1:10 (w/v). Two‐leaf‐stage seedlings were mechanically inoculated on the second true leaf using virus‐laden sap with carborundum abrasion.

Phylogenetic analysis

Orthologs of ZmCDPKs from Zea mays, Triticum aestivum, and Oryza sativa were utilized to construct a phylogenetic tree via the Construct/Test Neighbor‐Joining method in MEGA 7.0, incorporating 1,000 bootstrap replicates.

RNA extraction and RT‐qPCR

Total RNA was extracted from maize leaves using Trizol Plus (TaKaRa, Dalian, China), and subsequently treated with a gDNA wipe enzyme (Vazyme, Nanjing, China) to eliminate genomic DNA. The first‐strand cDNA was synthesized from 500 ng total RNA using HiScript III Q RT SuperMix kit (Vazyme, Nanjing, China) in a total volume of 10 μL following the manufacturer's instructions. Subsequently, 1.0 μL of diluted cDNA was used for RT‐qPCR analysis in a total volume of 10 μL with ChamQ SYBR qPCR Master Mix (Vazyme, Nanjing, China) on a PCR machine (Thermo Fisher, USA). Primers for RT‐qPCR were designed using an online tool (https://www.genscript.com.cn/tools/real-time-pcr-taqman-primer-design-tool). Relative gene expression was calculated using the 2^−ΔΔCT method as previously described, with ZmUBI expression serving as an internal control. The experiments were conducted in triplicate.

Yeast two‐hybrid assay

Maize leaves were collected and utilized for cDNA library construction following the manufacturer's protocol (Clontech, Beijing, China). ZmCDPK7 interacting host factors were screened according to the handbook provided by the Matchmaker Gold Yeast Two‐Hybrid System (Clontech, Beijing, China) (Ge et al., 2025).

For the Y2H assay, the Matchmaker Gold Y2H System was employed according to the manufacturer's instructions (Clontech, Beijing, China). Yeast cells (strain Y2H Gold) were co‐transformed with specific bait and prey constructs. All transformants were cultivated on a selective medium lacking tryptophan and leucine (SD/−TL) and on a high‐stringency selective medium devoid of tryptophan, leucine, histidine, and adenine (SD/−TLHA) to analyze potential interactions.

BiFC assay

The corresponding plasmids were individually transformed into the Agrobacterium strain GV3101. The Agrobacterium strains were co‐infiltrated at an OD₆₀₀ of 0.6 into the leaves of 4‐week‐old N. benthamiana in various combinations. Samples were observed using a NIKON A1HD25 laser scanning microscope at 48 hpi. YFP was excited at 514 nm and captured at 565–585 nm. Three independent experiments were conducted.

Co‐IP assay

Co‐immunoprecipitation assays were performed on protein extracts from N. benthamiana leaves. At 48 h post‐agroinfiltration, fluorescence signals were observed with Zeiss LSM710, and total proteins were extracted using lysis buffer (1 M Tris, pH 7.5, 5 M NaCl, 20% [v/v] Triton X‐100, 50% glycerol, 1 mM phenylmethylsulfonyl fluoride). For Co‐IP assays, 15 μL of agarose beads (GenScript, Nanjing, China) was incubated with 2 ml of crude protein at 4°C for 10 h and collected. Total cellular protein was extracted from fresh leaves in lysis buffer. The beads were collected, washed five times with a washing buffer, and analyzed by western blotting. Dilutions of HA, Flag, Myc, and GFP antibodies were 1:5,000. Blotted membranes were thoroughly washed and visualized using a chemiluminescence detection system (YESEN, Shanghai, China).

VIGS assay

The CMV–VIGS assay in maize was performed as previously reported (Zhou et al., 2018). Briefly, Agrobacterium (C58C1) cultures carrying pCMV201#2B_N81‐ZmLDAP2 or pCMV201#2B_N81‐GFP were co‐infiltrated with pCMV101 and pCMV301 into N. benthamiana leaves. At 3–4 dpi, infiltrated leaves were harvested, and crude extracts from these leaves were inoculated on maize seeds (inbred line B73). The inoculated seeds were kept at 25°C in the dark for 2 d, then transferred into pots with soil and grown in a growth chamber (20°C, 16 h of light and 18°C, 8 h of darkness). These plants were subjected to MCMV inoculation at the two‐leaf stage.

Western blot analysis

Soluble proteins from Nicotiana benthamiana were extracted using lysis buffer (1.0 M Tris‐HCl pH 6.8, 10% SDS, ddH₂O, β‐mercaptoethanol). The crude extracts were centrifuged at 12,000 g for 10 min. Proteins in the extracts were separated on 12% SDS‐PAGE and transferred onto nitrocellulose membranes. Antibodies or antiserum dilutions were prepared at 1:5,000. Hybridization signals were detected using an enhanced chemiluminescence system (eECL Western Blot Kit) (YEASEN, Shanghai, China).

Phosphorylation assays

The in vitro phosphorylation reactions were conducted in 30 μL of kinase buffer (25 mM Tris‐HCl, pH 7.5, 10 mM MgCl₂, 0.1 mM CaCl₂, 1 mM DTT, and 0.2 mM ATP) containing 1 μg ZmCDPK7‐His and 10 μg of substrate proteins at 28°C with gentle shaking for 0.5 h. For in planta phosphorylation reactions, proteins in the extracts were separated on 10% SDS‐PAGE and transferred onto nitrocellulose membranes. The phosphorylation of recombinant proteins from both in vitro and in planta phosphorylation reactions was analyzed by immunoblotting with an anti‐pSer/pThr antibody (Cat. no. ab117253, Abcam, UK) following protein separation by SDS‐PAGE.

Phosphorylation site prediction

To identify potential phosphorylation sites on the target proteins, bioinformatics approaches were employed in this study. Kinase‐specific phosphorylation sites within the protein sequences were predicted using the online tool GPS 5.0. Concurrently, supplementary predictions were generated using the NetPhos 3.1 server, integrating predictions from both tools.

Assays for APX activity, H₂O₂ concentration, and MDA content

APX activity: measured using the ascorbate peroxidase assay kit (Cat. no. BC0220, Solarbio, China). H₂O₂ concentration: quantified using the hydrogen peroxide assay kit (Cat. no. BC3590, Solarbio, China). MDA content: determined with the malondialdehyde assay kit (Cat. no. BC0025, Solarbio, China).

Electrolyte leakage assay

Ten leaf disks (1 cm² each) were excised with five biological replicates. Leaf tissues were briefly rinsed with deionized water to remove surface ions. Each replicate was floated in 10 mL of distilled water for 6 h at room temperature. Electrolyte leakage was measured using a conductivity meter (initial reading). The same solution containing the leaves was boiled at 90°C for 20 min. After cooling to room temperature, conductivity was remeasured (final reading). The percentage of ion leakage was calculated as follows: Percentage ion leakage = (initial conductivity/final conductivity) × 100%.

Histochemical staining of H₂O₂

The histochemical analysis of H₂O₂ using 3,3′‐DAB was performed as previously described (Yang et al., 2021). Maize leaf sections were vacuum‐infiltrated with 1 mg/mL DAB (Sigma, USA) in 50 mmol/L Tris‐acetate (pH 5.8) and then incubated at 25°C in darkness for 16 h. The staining solution was replaced with 90% (v/v) ethanol before the samples were heated. This process was repeated 4–5 times until the chlorophyll was completely removed.

Fusarium verticillioides inoculation

The fungal inoculation was performed following previously reported methods (Cao et al., 2023). Maize seeds were surface‐sterilized by soaking in 100 mL of 70% Ethanol (BP82031GAL, Fisher Scientific) for 3 min, followed by incubation in 500 mL of 50% commercial bleach (6% active sodium hypochlorite, Cat. NC9724348, Essendant) with Tween‐20 (BP337, Fisher) for 15 min. After five washes with sterile water and blot drying, five seeds were placed between wetted germination paper sheets (TopTech), rolled into a cylinder secured with rubber bands, and covered with aluminum foil (Cat. No. 01‐213‐105, Fisher Scientific). Each cylinder was placed in a glass tube containing an 800‐fold dilution of Peters Pro 20–20–20 water‐soluble fertilizer (ICL specialty fertilizers), reaching one‐fourth the height of the paper, and incubated for 6 d in a growth chamber (16 h light/27°C, 8 h dark/24°C, 50% humidity). On Day 8, healthy seedlings were transferred onto 1/2MS media (M5524, Sigma) in a 24 × 24 cm culture dish (BioSharp Life Sci). Root tips were inoculated with 20 μL of a hyphal mat homogenate (five plates blended to 125 mL, adjusted to 200 mL with ddH₂O), while mock inoculations used PDA plugs. The roots were covered with germination paper that was wetted with liquid 1/2MS media and then wrapped in aluminum foil. The covered dish was incubated for 48 h under the same growth conditions.

The disease resistance was tested on detached coleoptiles of wild‐type and knockout mutant maize plants. A single 5 mm² block of Fv PDA culture was placed on each leaf, which was removed 48 hpi. Infection was assessed based on the extent of discoloration and colonization 4 dpi at 25°C in the dark.

Drought stress treatment

Wild‐type and transgenic maize plants were subjected to drought stress by withholding water for 6 d at the four‐leaf stage. Their phenotypes were observed and photographed. On the fourth day of drought treatment, APX activity, H₂O₂ concentration, and MDA content were determined.

Statistical analysis

The data presented are means derived from three independent experiments. Statistical analysis was conducted using the Student's t‐test. Significance levels are indicated by asterisks, with P < 0.05 denoted by a single asterisk and P < 0.01 indicated by double asterisks. The intensity of the bands in western blotting was quantified using ImageJ software.

CONFLICTS OF INTEREST

The authors declare no conflicts of interest.

AUTHOR CONTRIBUTIONS

Y.S. and X.Y. designed the experiments. Y.Z., Q.W., Y.S., and Y.S. conducted the experiments. Y.Z., Q.W., X.H., N.L., H.L., and L.C. analyzed the data. Y.S. and X.Y. wrote the manuscript. Y.S., X.Y., H.Y., Z.F., and X.F. reviewed and edited the manuscript. All authors have read and approved the contents of this paper.

Supporting information

Additional Supporting Information may be found online in the supporting information tab for this article: http://onlinelibrary.wiley.com/doi/10.1111/jipb.70285/suppinfo

Figure S1. Identification of ZmCDPK7 in response to various stimuli

Figure S2. ZmCDPK7 positively regulates maize resistance to MCMV

Figure S3. ROS production was detected in MCMV infected Zmcdpk7 KO1 and Zmcdpk7 KO2 lines at 7 dpi

Figure S4. ZmCDPK7^K163R mutant failed to phosphorylate ZmAPX1 in vitro

Figure S5. The effect of MCMV infection on the expression of ZmAPX1 and ROS production

Figure S6. Silencing of ZmAPX1 promoted MCMV infection

Figure S7. ZmAPX1 negatively regulates ROS accumulation during MCMV infection

Figure S8. ZmRACK1 enhances ZmCDPK1‐mediated ZmAPX1 activity

Figure S9. ROS production was detected in MCMV‐infected Zmrack1 KO1 and Zmrack1 KO2 lines at 7 dpi

Figure S10. Silencing of ZmRACK1 promoted MCMV infection

Figure S11. Yeast two‐hybrid analysis of the combinations of BD + MCMV‐encoded proteins, and AD + BD‐ZmCDPK7

Figure S12. BiFC assay showing no interaction between P31 and ZmAPX1

Figure S13. The proposed working model for ZmRACK1–ZmCDPK7–ZmAPX1 functional module regulating immunity to MCMV

Table S1. Primer sequence

Table S2. The results of Y2H screening using ZmCDPK7 as a bait

Biographies

Zhang, Y. , Li, N. , Wang, Q. , Sun, Y. , Shi, Y. , Yu, L. , Han, X. , Feng, X. , Fan, Z. , Chen, L. , et al. (2026). The ZmRACK1–ZmCDPK7–ZmAPX1 module regulates plant antiviral immunity. J. Integr. Plant Biol. 68: 3830–3845.

Data availability statement

The genes investigated in this study are identified by the following accession numbers: ZmCDPK7: Zm00001d051502, ZmAPX1: Zm00001eb013080, and ZmRACK1: Zm00001eb294990.