MDP25‐VDAC3 Complex Orchestrates Actin Remodeling and Mitochondrial Dynamics to Modulate Innate Immunity in Arabidopsis

ABSTRACT

Host actin remodeling is critical for plant defense against pathogen infection, yet its specific functions remain to be fully explored. This study characterizes the role of the actin cytoskeleton in regulating mitochondrial dynamics to enhance plant defense. We demonstrate that the perception of the bacterial flagellin epitope induces excessive mitochondrial elongation, which facilitates the exchange of mitochondrial components for the functional recovery of damaged mitochondria. This elongation also promotes ATP and mitochondrial ROS production. The formation of elongated mitochondria largely depends on actin bundles in the cortical array. The contribution of actin bundles involves providing structural support for mitochondrial fusion and preventing already elongated mitochondria from undergoing fission. We further identified the interaction between the actin‐regulating protein MDP25 and the mitochondrial outer membrane protein VDAC3 in actin remodeling and mediating actin‐mitochondria interaction essential for mitochondrial fusion and elongation. Collectively, this study presents a mechanistic model for actin‐dependent mitochondrial elongation and its significance for plant immunity.

This study reveals that actin remodeling regulates mitochondrial elongation as a defense strategy in plants. Flagellin perception induces fusion‐dependent elongation, supported by cortical actin bundles, to repair damage and boost ATP and ROS production. The MDP25‐VDAC3 interaction links actin to mitochondria, establishing a model of actin‐dependent mitochondrial dynamics in immunity.

1. Introduction

The innate immune system plays a crucial role as the frontline defense against infections. In plant cells, the first layer of immunity involves the recognition of invading pathogens by cell surface receptors specifically evolved to detect a diverse range of microbe‐associated molecular patterns (MAMPs). Upon recognition, downstream defense responses are activated, including the production of reactive oxygen species (ROS), activation of mitogen‐activated protein kinase (MAPK) cascades, transcriptional reprogramming, apoplastic callose deposition, and stomatal closure [1].

Mitochondria are dynamic organelles responsible for ATP production, and their involvement in plant‐microbe interaction has also been reported [2]. For instance, fungal penetration induces the recruitment and immobilization of mitochondria at sites of attempted invasion in plant cells. Mitochondria arrested at the infection sites exhibit a redox imbalance, which may produce intracellular signals for defense activation [3, 4]. Several mitochondrial proteins, such as succinate dehydrogenase 1 (SDH1) [5], pre‐sequence translocase‐associated protein import motor (PAM16) [6], outer membrane 66 (OM66) [7] and oxidation resistance 2 (OXR2) [8], are implicated in plant resistance to pathogen infection. A recent study highlights the role of mitochondrial RNA processing mediated by Resistance To Phytophthora parasitica 7 (RTP7) in Arabidopsis host immunity [9]. Loss of RTP7 leads to increased mitochondrial ROS (mROS) and overall ROS production, resulting in enhanced plant resistance to a broad spectrum of pathogens [9]. Plant pathogens, in turn, have evolved strategies to target host cell mitochondria for their survival. For example, the rice fungal effector Avr‐Pita suppresses innate immunity by enhancing COX activity in rice mitochondria [10]. Additionally, the Magnaporthe oryzae effector MoCDIP4 inhibits rice immunity by targeting the mitochondria‐associated DjA9‐DRP1E protein complex to induce mitochondria fission [11], thus providing a functional correlation between mitochondrial dynamics and plant immunity.

The remodeling of the host actin cytoskeleton is a prominent response during the plant‐microbe interaction [12]. Upon the brief perception of MAMPs, the abundance of actin filaments in the cortical array increases in epidermal pavement cells [13, 14]. Disruption of the host actin cytoskeleton causes defects in surface immune receptor complex assembly, ROS production, callose deposition, activation of defense genes, and stomatal immunity. Consequently, this compromises the plant's resistance to pathogen infection [13, 14, 15, 16, 17, 18]. However, the molecular mechanisms underlying MAMP‐induced actin remodeling and its precise functions require further investigation. It is well established that the actin cytoskeleton regulates mitochondrial function and dynamics. While significant progress has been made in animal systems [19], less is known about the role of actin in mitochondrial biology in plants. Early studies have demonstrated that actin filaments are critical for mitochondrial distribution and movement [20, 21, 22, 23]. Treatments with actin‐disrupting agents or genetic inhibition of actin polymerization lead to reduced mitochondrial mobility [24, 25]. Actin filaments have also been implicated in maintaining mitochondrial membrane potential and Ca²⁺ homeostasis, which are crucial for root hair development and plant salt tolerance [25, 26]. Despite these findings, progress in understanding the functional connection between the actin cytoskeleton and mitochondria in plants has been limited over the past decade. Moreover, whether MAMP‐induced actin remodeling regulates mitochondrial dynamics and the biological significance of this interaction in plant disease resistance remains to be elucidated.

Microtubule‐Destabilizing Protein 25 (MDP25) in Arabidopsis was initially identified as a microtubule‐destabilizing and calcium binding protein involved in hypocotyl cell elongation [27, 28]. MDP25 is predominately localized to the plasma membrane (PM), and its function on cortical microtubules (MTs) is regulated by cytosolic calcium levels [28]. Subsequent research revealed that MDP25 is also an actin‐binding protein. It binds to and severs actin filaments, which is essential for pollen tube growth [29]. MDP25 expression is significantly upregulated by treatment with bacterial flagellin peptide mimic flg22 [30]. Moreover, MDP25 is involved in flg22‐induced resistance to fungal infection [31]. These observations prompted us to explore the role of MDP25 in regulating plant immunity. In the current investigation, we demonstrate that MDP25 interacts with the outer mitochondrial membrane (OMM) protein voltage‐dependent anion channel 3 (VDAC3) to regulate mitochondrial dynamics during plant innate immunity. Following flg22 treatment, MDP25‐VDAC3 complex regulates actin remodeling and actin‐mitochondria interaction, facilitating fusion‐dependent mitochondrial elongation. The changes in mitochondrial dynamics enhance the exchange of mitochondrial components for the functional recovery of damaged mitochondria, as well as the production of ATP and mROS for effective immune responses. In summary, our study uncovers the underlying molecular mechanism and the biological importance of host actin rearrangement in modulating mitochondrial dynamics during plant immunity.

2. Results

2.1. MDP25 Plays a Positive Role in Plant Immunity

MDP25 has previously been implicated in the regulation of flg22‐induced plant immunity [31]. However, the precise function of MDP25 in this process remains unclear. To address this, we examined multiple defense responses triggered by flg22 in WT plants, mdp25 mutants, and complementation lines expressing MDP25‐GFP under the control of its native promoter. These complementation lines have been characterized in previous studies and fully restore the developmental defects of the mdp25 mutant [28, 29, 32]. Flg22‐induced ROS production and MPK3/6/4 phosphorylation levels were comparable between WT and mdp25 mutants (Figure S1a,b). However, the loss of MDP25 resulted in significantly reduced callose deposition in the apoplast, and mutant stomata failed to close as effectively as WT plants following flg22 treatment (Figure S1c–f) [31]. In addition, a bacterial growth assay using the bacterial pathogen Pseudomonas syringae pv. tomato (Pst) DC3000 revealed that mdp25 plants supported significantly higher levels of bacteria compared to WT plants (Figure S1g). MDP25‐GFP complementation lines restored the mdp25 mutant phenotypes (Figure S1c–g). These data suggested that MDP25 positively regulates plant disease resistance.

Previous studies have shown that MDP25 can destabilize microtubules and sever actin filaments [28, 29]. Within the MDP25 amino acid sequence, Lys7 and Lys18 are critical sites for microtubule destabilization, while Asn5 is responsible for its actin binding and severing activity [28, 29]. To assess whether mutations in these key amino acids affect plant immunity, we conducted a bacterial growth assay on various mdp25 mutant plants expressing MDP25, MDP25 K7A, MDP25 K18A or MDP25 N5A driven by the endogenous promoter [28, 29]. As shown in Figure S1g, mutations in MDP25 K7 and MDP25 K18 restored the mdp25 mutant phenotype to the WT level, whereas the MDP25 N5A mutation did not (Figure S1g). These findings indicated that actin‐severing activity of MDP25 is crucial for its function in plant immunity.

2.2. MDP25 Interacts With VDACs

To elucidate how MDP25 regulates plant immunity, we aimed to identify MDP25‐interacting proteins. A GFP pull‐down assay was performed using MDP25‐GFP complementation lines or free GFP as a control. The immunoprecipitates of MDP25‐GFP and free GFP were analyzed by liquid chromatography‐tandem mass spectrometry (LC‐MS/MS)‐based proteomic analysis. We identified 128 proteins that were specifically enriched in the MDP25‐GFP pull‐down sample (Figure 1a; Data S1). Gene Ontology (GO) cellular component annotations revealed that many of these proteins were localized to the mitochondria (Figure 1b; Table S1). Among these mitochondrial proteins, VDAC1 caught our attention. VDACs are major mitochondrial outer membrane proteins that regulate the transport of metabolites between mitochondria and the cytoplasm. There are four VDAC genes (VDAC1‐4) expressed in Arabidopsis [33]. These proteins have been shown to maintain mitochondrial membrane potential and regulate hydrogen peroxide generation during plant disease resistance [34]. Given the functional connection between the cytoskeleton and mitochondria, we investigated whether MDP25 interacts with VDAC1. Using a split‐luciferase complementation assay in N. benthaminana, we found that MDP25 not only interacted with VDAC1 but also with other VDACs. Among these, VDAC3 exhibited the strongest binding with MDP25 (Figure 1c; Figure S2a). In contrast, no interaction was detected between MDP25 and other OMM proteins, such as OM66, ELM1, OM47 and TOM40 (Figure S2b) [7, 35, 36, 37]. We also noted that several matrix proteins were detected in the MDP25‐GFP pull‐down sample (Table S1). To assess their biological relevance, we tested the interaction between MDP25 and one representative matrix protein mtHSC70‐1 [38] and detected no binding (Figure S2b). This supports the possibility that their recovery reflects loss of mitochondrial compartmental integrity during protein extraction rather than genuine in vivo interactions.

FIGURE 1.

MDP25 interacts with VDAC3. (a) Potential MDP25‐interacting proteins enriched in the MDP25‐GFP pulldown sample. (b) Gene Ontology cellular component annotations are analyzed; this analysis identified 22 mitochondrial proteins that may interact with MDP25. (c) Luciferase complementation imaging assay suggests an interaction between MDP25 and VDAC3 in planta. nLUC, N‐terminal fragment of firefly luciferase; cLUC, C‐terminal fragment of firefly luciferase. (d) In vitro protein pull‐down assay demonstrates the direct interaction of MDP25 and VDAC3. Purified, recombinant His‐tagged VDAC3 cosedimented with purified MDP25 fused to a GST tag (MDP25‐GST), but not with purified GST alone. (e) Co‐IP assay was performed in N. benthamiana leaves co‐transformed with the indicated constructs. Total proteins (input) were subjected to immunoprecipitation with anti‐GFP beads followed by immunoblotting analysis. (f‐g) flg22‐induced Ca²⁺ influx promotes MDP25 redistribution from the PM to the cytosol and facilitates interaction with VDAC3. (f) Immunoblot analysis of MDP25‐GFP in cytosolic (Cyt) and PM‐enriched fractions from seedlings treated with mock, flg22, or flg22 with LaCl₃. MDP25‐GFP was detected by anti‐GFP antibodies. PM H⁺‐ATPase and Rubisco serve as PM and cytosolic markers, respectively. (g) Quantification of MDP25‐GFP levels in cytosolic and PM fractions normalized to total signal. Band intensity was quantified by densitometric analyses. Data were quantified from four independent biological replicates. (h‐i) FRET‐FLIM analysis of the interaction between MDP25 and VDAC3. (h) Intensity and fluorescence lifetime maps of VDAC3‐GFP when expressed alone or co‐expressed with MDP25‐mCherry. Scale bar=2 µm. Scale varies from the shortest lifetime of 1.6 ns to the longest lifetime of 2.6 ns. (i) Quantification of the average fluorescence lifetime of VDAC3‐GFP when expressed alone or co‐expressed with MDP25‐mCherry. Seven‐day‐old seedlings were treated with mock, 10 µM flg22 with or without 10 mM LaCl₃ for 3 h before protein extraction or FRET‐FLIM analyses. (j) Both MDP25 and VDAC3 are required for plant resistance to DC3000 infection. Two‐week‐old seedlings of indicated genotypes were spray inoculated with DC3000. Bacterial growth was measured at 2 d post infection (see methods for experimental details). Values are means ± SD. N in (j)=10 biologically independent samples. Experiments were repeated at least three times with similar results. Significance in (g) was assessed by two‐way ANOVA with Tukey's multiple comparisons test, and in (i) and (j) by one‐way ANOVA with Tukey's multiple comparisons test. Different letters indicate significant differences at P<0.05.

Based on the stronger interaction between VDAC3 with MDP25, we focused on VDAC3 in this study. In a pull‐down assay, His‐tagged VDAC3 was able to bind GST‐MDP25 but not GST alone, indicating that MDP25 can directly interact with VDAC3 in vitro (Figure 1d). Moreover, a coimmunoprecipitation (Co‐IP) assay further confirmed the MDP25/VDAC3 interaction (Figure 1e). MDP25 is predominantly localized to the PM [28], raising the question of how it could interact with mitochondrial OMM proteins in the cytosol. Previous studies have shown that elevated cytosolic Ca²⁺ levels under salt stress can trigger partial dissociation of MDP25 from the PM [39]. To test whether flg22 elicits a similar redistribution, we fractionated cytosolic and PM components from flg22‐treated leaves and performed immunoblotting. As expected, MDP25‐GFP was predominantly detected in the PM fraction, but cytosolic signals were also evident (Figure 1f,g). Following flg22 treatment, MDP25‐GFP levels increased in the cytosol (Figure 1f,g; Figure S2c,d). We also detected the overlapping signals between cytosolic MDP25‐GFP and mitochondria (Figure S2e). The increase in cytosolic MDP25‐GFP levels was blocked by the calcium channel blocker LaCl₃, indicating that flg22‐triggered Ca²⁺ influx mediates MDP25 translocation from the PM to the cytosol (Figure 1f,g). We further employed fluorescence resonance energy transfer (FRET)‐fluorescence lifetime imaging microscopy (FLIM) assay to directly visualize MDP25‐VDAC3 interaction in living cells. The average fluorescence lifetime of VDAC3‐GFP was significantly reduced in the presence of MDP25‐mCherry under mock conditions, suggesting a direct interaction. This reduction was further enhanced by flg22 treatment and abolished by LaCl₃ (Figure 1h,i). Taken together, these results demonstrate that flg22 induces Ca²⁺‐dependent dissociation of MDP25 from the PM into the cytosol, where it directly interacts with VDAC3 on mitochondria.

Next, we assessed various immune responses in a VDAC3 knockdown mutant (vdac3‐1) [40] (Figure S2f). Similar to the mdp25 mutant, reduced VDAC3 levels in vdac3‐1 led to significantly impaired callose deposition and stomatal closure following flg22 treatment (Figure S2g–j). In addition, vdac3‐1 mutants exhibited increased susceptibility to DC3000 infection compared to WT plants (Figure S2k). The defects in bacterial growth observed in vdac3‐1 mutants were restored by expressing VDAC3 under the control of the U10 promoter in the vdac3‐1 background (VDAC3‐OE; Figure S2f,k). We also generated a double mutant by crossing the homozygous single mutants (mdp25vdac3‐1). As shown in Figure 1j, the mdp25vdac3‐1 double mutant exhibited slightly more severe bacterial growth phenotypes compared to the single mutants, although the differences were not statistically significant (Figure 1j). Overall, our findings underscore the positive roles of both MDP25 and VDAC3 in plant immunity.

2.3. MDP25 and VDAC3 Are Required for flg22‐Induced Mitochondrial Elongation

That MDP25 interacts with VDAC3 indicates its involvement in mitochondrial dynamics. To explore this possibility, we initially investigated whether flg22 treatment alters mitochondrial morphology. Confocal microscopy was employed to visualize mitochondria in cotyledon pavement cells expressing mitochondrial matrix‐targeted GFP (mt‐GFP). In untreated cells, mitochondria predominantly exhibited spherical to ovoid structures (Figure 2a) [4]. Following flg22 treatment, mitochondria underwent considerable elongation, adopting a tubular shape (Figure 2a). To further scrutinize mitochondrial morphology at the electron microscopic level, ultrathin sections were prepared from cotyledons of WT plants treated with mock or flg22. Transmission electron microscopy (TEM) unveiled abnormal elongation of mitochondria in flg22‐treated plants compared to mock controls (Figure S3a), validating the morphological changes observed via confocal microscopy. Notably, the internal structures of mitochondria, including the matrix and cristae, exhibited no discernible differences between spherical or elongated mitochondria (Figure S3a).

FIGURE 2.

MDP25 and VDAC3 regulate fusion‐dependent mitochondrial elongation triggered by flg22. (a) Representative images showing mitochondrial morphology in WT, mdp25, vdac3‐1 single and mdp25vdac3‐1 double mutant after mock or 10 µM flg22 treatment for 12 h. Images were taken from cotyledon pavement cells of 2‐week‐old seedlings expressing mt‐GFP. Scale bar=2 µm. (b) The ratio between length and width of mitochondria was quantified in the indicated genotypes. (c) The mitochondrial length was analyzed and categorized into three populations: spherical (0–2 µm), intermediate (2–4 µm) and elongated (>4 µm). The frequency of each population within the total mitochondria quantified was calculated in WT and mutants after mock and flg22 treatment. (d,e) The rates of mitochondrial fusion (d) and fission (e), expressed as total events per mitochondrion per minute, were analyzed in WT and mutants. (f) The fusion/fission ratio was quantified by dividing the rate of fusion by the rate of fission in each cell. Images were collected and analyzed from cotyledon pavement cells of 2‐week‐old seedlings after mock or 10 µM flg22 treatment for 12 h. Values are means ± SD. The data in (b) and (c) were quantified from the same set of confocal images presented in (a). All mitochondria within each cell were analyzed, with approximately 30 cells included in the quantification. N in (b) denotes the number of cells. N in (c) denotes the number of mitochondria from 30 cells. N in (d–f) represents the number of time‐lapse images. Results in (f) are quantified from the same images in (d,e). Experiments were repeated at least three times with similar results. Different letters indicate significant differences at P<0.05, as determined by two‐way ANOVA with Tukey's multiple comparisons test.

The alterations in morphology were further quantified by assessing the ratio of mitochondrial length to width from confocal images. Following flg22 treatment, the length/width ratio significantly increased compared to the mock treatment (Figure S3b). Similar to flg22, treatment with another bacterial MAMP, elf18, or a fungal MAMP, chitin, also led to mitochondrial elongation. As a negative control, we applied treatments with flgII‐28, a flagellin epitope perceived by few species belonging to the Solanaceae family [41, 42]. No obvious changes in mitochondrial morphology were detected in response to flgII‐28 (Figure S3b). These data suggested that mitochondrial elongation is a common cellular response to different MAMPs. The flg22‐induced increase in the length/width ratio primarily resulted from changes in mitochondrial length, not width (Figure S3c). Throughout a time‐course treatment with flg22, the length/width ratio increased at 6 h post‐elicitation, reached its peak at 12 h, then gradually decreased from 24 h onward, eventually returning to a level comparable to that at 0 h (Figure S3d). In the flagellin receptor mutant fls2 [43], the length/width ratio did not increase as in WT plants following flg22 treatment (Figure S3e), indicating that changes in mitochondrial morphology are dependent on flg22 perception by its cognate receptor.

To investigate whether MDP25 and VDAC3 are involved in flg22‐induced mitochondrial elongation, we conducted live‐cell imaging in WT, mdp25, vdac3‐1 single and mdp25vdac3‐1 double mutant cells expressing mt‐GFP. Under mock conditions, the length/width ratios of mitochondria in mdp25, vdac3‐1 single and double mutant cells did not significantly differ from those in WT cells. In response to flg22, all mutants failed to elongate mitochondria to the same extent as WT (Figure 2a,b). In addition, the mdp25vdac3‐1 double mutants did not exhibit an additive phenotype compared to the single mutants. (Figure 2a,b). We further analyzed the length/width ratio in various MDP25 complementation lines. The mdp25 mutant plants expressing MDP25 K7A or K18A showed WT mitochondrial responses to flg22 treatment (Figure S4a,b). MDP25 N5A expression failed to rescue the mitochondrial phenotypes in the mdp25 mutant (Figure S4a,b). Expression of VDAC3 in vdac3‐1 mutants restored their mitochondrial responses to flg22 (Figure S4c). The mitochondrial volume was quantified in WT and different mutants. Under resting conditions, the mitochondria in WT cells had an average volume of 0.25 µm³, the volume of mitochondria in mdp25 cells showed no significant differences from WT, whereas mitochondria in vdac3‐1 cells were significantly larger than those in WT and mdp25 cells (Figure S4d). After flg22 treatment, the volume of WT mitochondria increased compared to the mock control. The mitochondria volume in mdp25 also increased but to a less extent than WT. By contrast, the mitochondria sizes did not change in vdac3‐1 single and mdp25vdac3‐1 double mutant after flg22 treatment compared to mock (Figure S4d).

Based on measurements of mitochondrial length, we categorized mitochondrial morphology into three classes: spherical (<2 µm), intermediate (2–4 µm), and elongated (>4 µm). The distribution of each class within the total mitochondrial population was analyzed. In mock‐treated cells, mitochondrial morphology appeared relatively uniform, with approximately 90% of the population displaying a spherical shape and 10% featuring intermediate mitochondria (Figure 2c). In the stimulated group, however, mitochondrial morphology became more diverse. The number of spherical mitochondria decreased, while tubular mitochondria increased. The majority (>60%) of mitochondrial lengths exceeded 2 µm, with approximately 25% surpassing 4 µm (Figure 2c). Mutations in MDP25 or VDAC3 led to a significant reduction in the population of intermediate and elongated mitochondria after flg22 treatment, particularly in the formation of mitochondria longer than 4 µm (Figure 2c). These results indicated the essential role of both MDP25 and VDAC3 in flg22‐triggered mitochondrial elongation. Moreover, the rise in the proportion of elongated mitochondria coincided with a decrease in the number of mitochondria in each cell (Figure S4f), suggesting that the flg22‐induced mitochondrial elongation is likely due to the fusion/fission imbalance.

To further demonstrate whether the compromised disease resistance in mdp25 and vdac3‐1 mutants is a result from the impaired mitochondrial elongation, we applied treatments with the mitochondrial division inhibitor (MIDI) [44]. Treatments with MIDI sufficiently led to elongated mitochondria in WT, mdp25 and vdac3‐1 mutants (Figure S5a,b). Importantly, the enhanced susceptibility to bacterial infection in both mutants can be partially suppressed by MIDI treatments (Figure S5c). These results suggested that MDP25 and VDAC3 contribute to plant disease resistance, at least in part, through mitochondrial elongation.

2.4. MDP25 and VDAC3 Regulate Fusion‐Dependent Mitochondrial Elongation Triggered by flg22

To gain a deeper understanding of the mechanism underlying flg22‐induced mitochondrial elongation, we conducted time‐lapse imaging to capture mitochondrial fusion and fission after mock or flg22 treatment. It's worth noting that the brief intramitochondrial association may lead to transient fusion followed by fission occurring at or near the fusion site, a phenomenon referred to as “kiss and run” [45, 46]. In this study, we specifically focused on complete fusion, where interacting mitochondria give rise to a single organelle that later undergoes fission at different sites [46]. Having observed 127 instances of fusion events in WT cells, we identified two forms of mitochondrial mergers. Ninety percent of the fusions occurred through longitudinal merging along the mitochondria's long axis (Figure S6a,c; Movie S1). Additionally, mitochondrial fusion could also initiate from oblique or lateral interactions, constituting 10% of the total fusion events (Figure S6b,c; Movie S2).

The rates of fusion and fission were quantified in WT and mutant cells subjected to mock and flg22 treatment. As depicted in Figure 2d–f, both fusion and fission events were infrequent in mock‐treated cells. Only 0.0067 or 0.0073 apparent fusion or fission events were observed per mitochondrion during 1‐min recordings in WT, respectively. No significant differences in fusion or fission rates were observed among WT, mdp25, vdac3‐1, and their respective complementation lines treated with mock (Figure 2d,e; Figure S7). Following flg22 treatment, both fusion and fission rates significantly increased in WT compared to the mock control, with fusion events occurring more frequently than fission (Figure 2d–f; Figure S7). In mdp25 and vdac3‐1 mutant cells treated with flg22, rates of fusion, but not fission, exhibited a significant decrease compared to WT (Figure 2d–f; Figure S7). Collectively, these results indicated that flg22‐induced mitochondrial elongation is a consequence of enhanced mitochondrial fusion, which requires the function of both MDP25 and VDAC3.

2.5. Mitochondrial Elongation Mediated by MDP25 and VDAC3 Facilitates the Functional Recovery of Damaged Mitochondria, and the Production of mROS and ATP in Response to flg22

To assess differences in the functional status of mitochondria in WT and mutants after mock and flg22 treatment, we examined the mitochondrial membrane potential using tetramethylrhodamine methyl ester (TMRM) staining. In the initial 2 h of flg22 treatment, the TMRM signals significantly decreased compared to 0 h, indicating compromised mitochondrial activity. The membrane potential fully recovered at 6 h and 12 h post‐flg22 treatment (Figure 3a,b), coinciding with the onset of mitochondrial elongation (Figure S3d). These data suggested that fusion‐mediated mitochondrial elongation likely contributes to the functional recovery of damaged mitochondria. Mitochondrial fusion has been shown to enable the mixing of mitochondrial content throughout the network to optimize function [47]. Therefore, the mitochondria‐targeted photoconvertible protein (mt‐Dendra2) was used to assay mitochondrial matrix fusion [48]. Matrix‐localized Dendra2 was photoconverted within a sub‐population of mitochondria in pavement cells. After photoconversion, the mitochondria were shifted to red fluorescence (Figure S8a,b). Time‐lapse imaging showed that fusion between red and green mitochondria led to the transfer of fluorescence signals, indicating matrix mixing (Figure S8a,b). The percentage reduction in numbers of green mitochondria, due to matrix contents exchange, was significantly greater at the end of the assay time period in flg22‐treated cells (36%) compared to mock‐treated cells (10%; Figure S8c,d). These data suggested that enhanced exchange of mitochondrial content via fusion is likely a mitochondrial quality control mechanism upon flg22 stimulation. Mitochondrial activity was further examined in mdp25, vdac3‐1 mutants through mitochondrial membrane potential analysis. Under resting conditions, both mutants exhibited reduced mitochondrial membrane potential, as indicated by lower TMRM fluorescence compared with WT. These defects were rescued in their respective complementation lines (Figure 3a,b; Figure S9a). Following flg22 elicitation, the decrease in TMRM fluorescence observed in WT at 0.5 and 2 h post‐treatment was not detected in mdp25 and vdac3‐1 mutants. Moreover, the recovery of membrane potential did not occur at 6 h after flg22 treatment in these mutants (Figure 3a,b).

FIGURE 3.

Mitochondrial elongation facilitates the functional recovery of damaged mitochondria, the production of mROS and ATP in response to flg22. (a) Representative images of mitochondria stained with TMRM in WT, mdp25 and vdac3‐1 mutant after mock and flg22 treatment for 0 h, 0.5 h and 12 h. Scale bar=2 µm. (b) Quantification of TMRM fluorescent intensity in WT, mdp25 and vdac3‐1 mutant cells at the indicated timepoints after flg22 treatment. (c) Representative images of mitochondria after MitoSOX Red staining in cotyledon pavement cells expressing mt‐GFP. Scale bar=2 µm. (d) Quantification of MitoSOX Red fluorescent intensity in the indicated genotypes. (e) Ratiometric imaging of ATP biosensor ATeam1.03‐nD/nA in cotyledon pavement cells after mock and flg22 treatment. Fluorescence of YFP and CFP was recorded, and the ratio was plotted as pseudo‐color images. Scale bar=20 µm. (f) Quantification of YFP:CFP ratio in cells treated with mock or flg22. Images in (c,e) were collected from cotyledon pavement cells of 2‐week‐old seedlings after mock or 10 µM flg22 treatment for 12 h and used for analyses in (d,f). Values are means ± SD. N in (b,d) denotes the number of cells from 7 seedlings for each genotype and treatment. N in (f) denotes the number of ROIs (73 µm × 36 µm) from more than 30 images and 7 seedlings for each genotype and treatment. Experiments were repeated at least three times with similar results. Different letters indicate significant differences at P<0.05, as determined by two‐way ANOVA with Tukey's multiple comparisons test.

The important role of mROS in plant immunity has been demonstrated [2, 9]. Thus, we analyzed whether mitochondrial elongation enhances mROS production by staining mitochondria with mROS‐specific fluorescent probe MitoSOX Red [9]. In unstimulated WT cells, only a few mitochondria were stained by MitoSOX Red, whereas more mitochondria showed stronger mROS signals in flg22‐treated cells (Figure 3c,d). However, flg22‐induced mROS production was compromised in the mdp25 and vdac3‐1 mutants, but fully restored in their respective complementation lines (Figure 3c,d; Figure S9b,c). To explore whether flg22‐induced mitochondrial elongation influences ATP generation, we measured ATP concentration in cells using the ratiometric ATP biosensor, ATeam1.03‐nD/nA. ATP binding induces a conformational change in the sensor structure, leading to increased FRET efficiency [49]. Cellular ATP levels significantly increased in WT cells post‐flg22 treatment compared to mock‐treated cells. Consistent with the diminished membrane potential, both mdp25 and vdac3‐1 mutant cells exhibited lower ATP levels than WT under resting conditions. Additionally, the increased ATP production in response to flg22 was hindered by mutations in either MDP25 or VDAC3 (Figure 3e,f). These findings suggested that mitochondrial elongation mediated by MDP25 and VDAC3 facilitates the functional recovery of damaged mitochondria, and supports the production of mROS and ATP during defense.

2.6. MDP25 and VDAC3 Are Required for flg22‐Induced Actin Remodeling

The fact that MDP25 N5A failed to rescue the mitochondrial responses to flg22 in the mdp25 mutant indicates the essential role of its actin‐severing activity. However, we first investigated which cytoskeletal element participates in flg22‐induced mitochondrial elongation. As shown in Figure S10a,b, the complete disruption of cortical actin or MT structures by LatB or oryzalin, respectively, resulted in impaired mitochondrial responses to flg22 (Figure S10a,b). However, the effects of LatB were more severe than oryzalin, suggesting the predominant contribution of the actin cytoskeleton in this process (Figure S10b).

To explore the relationship between actin dynamics and mitochondrial elongation, we assessed the changes in the actin organization at different time points after flg22 treatment. Consistent with previous results, treatments with flg22 for 0–3 h led to a more dense cortical actin array in cotyledon pavement cells compared to mock treatments [13, 14]. However, actin filaments started to form bundles when treated with flg22 longer than 3 h (Figure S10c). To verify these changes, we analyzed the optical densities of actin filament structures by measuring the percentage of occupancy of actin filaments [13]. To estimate the extent of actin bundling in the cortical array, we calculated the intensity of each filamentous structure within a cell and normalized these intensities to those of single actin filaments. Higher relative intensity levels indicate the formation of thicker actin bundles. When cells were treated with flg22 for 0 to 3 h, the densities of actin filaments increased, whereas the relative intensity levels of filamentous actin structures did not change compared to mock‐treated cells (Figure S10c–e). However, after prolonged flg22 treatment, the overall actin network appeared less dense between 12 and 24 h, but returned to a level comparable to the 0‐h baseline by 48 h (Figure S10c,d). Instead, robust formation of actin bundles in the cortical array was detected from 6 h onward in flg22‐treated cells, as indicated by the increase in the relative intensity levels of filamentous actin structures (Figure S10c,e). The extent of bundling reached its peak at 12 h post‐treatment, began to decline after 24 h and eventually returned to baseline levels by 48 h following flg22 exposure (Figure S10c,e). In contrast, MT organization was not significantly affected by flg22 (Figure S10f,g). These findings suggested that flg22 treatment for different time periods induces distinct actin responses. Short‐term treatment leads to increased actin density, while extended stimulation induces actin bundle formation in the cortical array of cotyledon pavement cells.

To investigate the underlying mechanisms of flg22‐induced actin rearrangement, we examined the dynamic behaviors of single filaments. As shown in Table S2, the increased actin density following flg22 treatment for 0–3 h resulted from enhanced filament nucleation and decreased filament disassembly (Table S2) [50]. After extended flg22 treatment, the frequency of filament nucleation did not differ significantly from the mock group. However, the severing frequency remained reduced, and filaments tended to form bundles, as indicated by an upregulated frequency of bundling (Table S2). Collectively, the results from single filament analysis demonstrated that different durations of flg22 treatment cause different dynamic behaviors of actin filaments.

Next, we assessed whether MDP25 and VDAC3 are required for flg22‐induced actin remodeling. Under mock conditions, actin arrays in mdp25 mutant cells exhibited reduced filament abundance compared to WT. However, the relative intensity levels of actin structures in mdp25 mutants were not significantly different from WT (Figure 4; Figure S11a–c). Furthermore, both flg22‐induced early increase in actin density and later bundle formation were abolished in the mdp25 mutant (Figure 4; Figure S11a–c). Compared to WT, loss of MDP25 did not alter the gross organization of the cortical MT array in cotyledon pavement cells. Moreover, MTs in mdp25 mutants did not respond to flg22 stimulation (Figure S10f,g). The actin defects in mdp25 mutants were restored by the expression of MDP25, MDP25 K7A and K18A, but not by MDP25 N5A (Figure S11d–f), suggesting that sites for microtubule destabilization are dispensable for MDP25 to regulate actin dynamics. Notably, MDP25, MDP25 K7A and MDP25 K18A were still able to promote mitochondrial elongation upon flg22 treatment. In contrast, MDP25 N5A failed to induce such mitochondrial changes (Figure S4a,b). This pattern closely mirrors their corresponding effects on actin filament organization, indicating that the ability of MDP25 to remodel the actin cytoskeleton is linked to its role in coordinating mitochondrial responses. Similarly, the vdac3‐1 single mutant cells displayed similar actin organization and response to flg22 as observed in the mdp25 mutant, despite VDAC3 not being previously associated with actin‐regulating activity (Figure 4; Figure S11a–c). The actin phenotypes in mdp25vdac3‐1 double mutant cells were comparable to the single mutants (Figure 4; Figure S11a–c). Collectively, these findings demonstrated that both MDP25 and VDAC3 are involved in actin rearrangements in response to flg22.

FIGURE 4.

MDP25 and VDAC3 are required for the formation of actin bundles induced by flg22. (a) Representative confocal images showing actin organization in the indicated genotypes after mock or flg22 treatment. Scale bar=2 µm. (b,c) The density (b) and relative intensity level (c) of filamentous actin structures measured in the indicated genotypes after mock or flg22 treatment. Images were collected and analyzed from cotyledon pavement cells expressing GFP‐fABD2. Two‐week‐old seedlings of the indicate genotypes were treated with mock or 10 µM flg22 for 12 h. Values are means ± SD. N in (b,c) indicates the number of images from 10 seedlings for each genotype and treatment. Experiments were repeated at least three times with similar results. Different letters indicate significant differences at P<0.05, as determined by two‐way ANOVA with Tukey's multiple comparisons test.

2.7. MDP25 and VDAC3 Mediate Actin‐Mitochondria Interaction in Response to flg22

The temporal correlation between actin bundle formation and mitochondrial dynamics suggests a functional link between these two cellular processes (Figure S10c–e; Figure S3d). To test this, dual‐color imaging was conducted on plants labeled with GFP‐fABD2 and mt‐mScarlet in WT and mutant lines. In mock‐treated cells, spherical mitochondria displayed localized oscillatory movements and transiently interacted with surrounding actin filaments. They also traveled along actin tracks (Figure 5a–c; Movie S3). Consistent with previous observations (Figure 2d–f), both fusion and fission events were rarely detected. Upon flg22 treatment, more mitochondria remained attached to actin bundles for extended periods, and frequent end‐to‐end fusion were observed along bundled actin filaments (Figure 5a; Movie S4). We found that three distinct mitochondrial morphology classes exhibited varying tendencies to associate with actin bundles. Among spherical mitochondria, only 30% of this subset interacted with actin bundles. In contrast, nearly all elongated mitochondria were attached to and moved along actin bundles (Figure 5a; Figure S12a; Movies S3 and S4). We also categorized all fusion events monitored into two groups based on the resulting mitochondrial length after fusion (<4 µm or >4 µm). From each category, we quantified the percentage of fusion events occurring on or off actin bundles. Approximately 60% of fusion events forming mitochondria shorter than 4 µm occurred on actin bundles, whereas all fusion events forming mitochondria longer than 4 µm occurred exclusively on actin bundles (Figure S12b). These findings suggested that mitochondrial fusion preferentially occurs on actin bundles in response to flg22. Elongated mitochondria, once detached from bundled actin filaments, tended to undergo fission at multiple sites (Figure S12c; Movie S5). Collectively, these results indicated that actin bundles contribute to mitochondrial elongation by providing structural support for mitochondrial fusion and preventing fission of elongated mitochondria.

FIGURE 5.

MDP25 and VDAC3 are required for actin‐mitochondria interaction in response to flg22. (a) Representative time‐lapse images showing dynamic interaction between mitochondria (green) and actin filaments (magenta) in WT, mdp25 and vdac3‐1 mutant after mock and flg22 treatment. Scale bars=2 µm. In WT cells treated with mock or mutant cells, spherical or intermediate mitochondria (indicated by arrowheads) displayed localized oscillatory movements and transiently interacted with surrounding actin filaments. Fusion events on actin bundles were rarely detected. Also see Movies S3 and S6–S9. In WT treated with flg22, more mitochondria remained attached to actin bundles for extended periods. Two mitochondria (indicated by white and yellow arrowheads) aligned and moved toward each other along this bundle. After these organelles met, end‐to‐end fusion took place, resulting in the formation of an elongated mitochondrion (indicated by the white arrowhead), also see Movie S4. The regions in the dashed line boxes are enlarged in the panels below. (b,c) Quantification of the number (b) and duration (c) of mitochondria associated with actin bundles in the indicated genotypes. (d) The frequency of fusion occurring on actin bundles was analyzed in the indicated genotypes after flg22 treatment. (e) Analysis of severing frequency of peri‐mitochondrial actin filaments in the indicated genotypes. Images were collected and analyzed from cotyledon pavement cells expressing GFP‐fABD2 and mt‐mScarlet. Two‐week‐old seedlings of the indicate genotypes were treated with mock or 10 µM flg22 for 12 h. Values are means ± SD. N in (b) denotes the number of ROIs (9 µm × 10 µm) from 23 cells and 7 seedlings for each genotype and treatment. N in (c,d) denotes the number of time‐lapse sequences from 7 seedlings. N in (e) represents the number of ROIs (10 µm x 10 µm) from more than 19 cells and 7 seedlings for each genotype and treatment. Experiments were repeated at least three times with similar results. Significance in (b), (c), and (e) was assessed by two‐way ANOVA with Tukey's multiple comparisons test, and in (d) by one‐way ANOVA with Tukey's multiple comparisons test. Different letters indicate significant differences at P<0.05.

The defects in flg22‐induced actin bundle formation observed in the mdp25 and vdac3‐1 mutants indicated a potential reduction in the availability of actin structures for mitochondrial attachment and fusion. To address this, we analyzed the number and duration of mitochondria associated with actin bundles in WT and mutants. Under mock conditions, no significant differences were detected between WT and mutants (Figure 5a–c; Movies S3, S6, and S8). Following flg22 treatment, fewer mitochondria attached to actin bundles in mdp25 and vdac3‐1 mutants compared to WT. Additionally, mitochondrial association time with actin bundles did not increase in response to flg22 treatment in these mutants (Figure 5a–c; Movies S4, S7, and S9). Moreover, unlike WT, the mitochondrial fusion on actin bundles were significantly reduced in mdp25 and vdac3‐1 mutants (Figure 5a,d; Movies S4, S7, and S9). These findings suggested that following flg22‐induced fusion initiation, fusion of small or short tubular mitochondria is less dependent on actin bundles. However, further mitochondrial elongation requires attachment to and fusion on actin bundles. In mdp25 or vdac3‐1 mutants, defects in actin bundle formation and mitochondria‐actin association result in a significant reduction in mitochondrial fusion on actin bundles, leading to a decreased formation of intermediate and elongated mitochondria following flg22 treatment (Figure 2c,d).

To further substantiate these observations, we next performed a quantitative analysis of mitochondria‐adjacent actin dynamics in WT and mutant cells. As shown in Figure 5e, flg22 treatment significantly reduced the severing frequency of peri‐mitochondrial actin filaments in WT cells. This suggested that localized actin remodeling is tightly regulated to provide stable filamentous structures that facilitate mitochondrial fusion. In the mdp25 mutant, basal severing activity was lower than in WT, in agreement with MDP25's role as a severing protein. Importantly, severing frequency of peri‐mitochondrial actin filaments in mdp25 mutants did not further decrease upon flg22 treatment. The vdac3‐1 mutant also failed to show reduced severing under flg22 conditions compared to WT (Figure 5e). Together, these results indicate that both MDP25 and VDAC3 are required for remodeling of mitochondria‐adjacent actin filaments.

2.8. MDP25/VDAC3 Interaction Is Important for Actin and Mitochondrial Responses to flg22

Although the mdp25 and vdac3‐1 mutants exhibit similar defects in actin organization and mitochondrial morphology, we cannot exclude the possibility that these two proteins function independently in these processes. To determine the biological relevance of MDP25/VDAC3 complex formation in actin remodeling and mitochondrial elongation, we employed AlphaFold2, a deep‐learning‐based protein structure prediction tool [51], to identify the specific residue(s) critical for the MDP25/VDAC3 interaction. AlphaFold 2 predicted that K80 in MDP25 and D168 in VDAC3 are key residues for this interaction (Figure 6a). To validate this prediction, we mutated MDP25 K80 and VDAC3 D168 to G. The VDAC3 D168 mutation inhibited the MDP25/VDAC3 interaction, whereas MDP25 K80 mutation did not (Figure 6b,c; Figure S13a). These data suggested that VDAC3 D168 is critical for MDP25/VDAC3 complex formation.

FIGURE 6.

MDP25/VDAC3 interaction is important for actin and mitochondrial responses to flg22. (a) Cartoon representations of the predicted interaction between MDP25 and VDAC3 highlighting K80 of MDP25 and D168 of VDAC3. (b,c) Luciferase complementation imaging assay (b) and in vitro protein pull‐down assay (c) showed that the interaction between MDP25 and VDAC3 was disrupted by the VDAC3 D168G mutation. (d) The ratio between the length and width of mitochondria was quantified in the indicated genotypes. (e,f) The rates of mitochondrial fusion (e) and fission (f) were quantified in the indicated genotypes. (g,h) The density (g) and relative intensity level (h) of filamentous actin structures measured in the indicated genotypes. Data were obtained from images taken from cotyledon pavement cells expressing mt‐GFP (d‐f) or GFP‐fABD2 (g,h). Two‐week‐old seedlings were treatment with mock or 10 µM flg22 for 12 h. (i) Bacterial growth was measured in WT, vdac3‐1 mutant, and vdac3‐1 mutant expressing VDAC3‐GFP or VDAC3^D168G‐GFP. Data in (d‐f) were obtained from WT, vdac3‐1 mutants, and VDAC3‐GFP or VDAC3^D168G‐GFP complementation lines after staining with 500 nM MitoTracker for 1 h. Data in (g,h) were obtained from WT, vdac3‐1 mutants, and VDAC3‐mCherry or VDAC3^D168G‐mCherry complementation lines expressing GFP‐fABD2. Values are means ± SD. N in (d) denotes the number of cells from 7 seedlings quantified for each genotype and treatment. N in (e,f) denotes represents the number of time‐lapse images from 7 seedlings quantified for each genotype and treatment. N in (g,h) indicates the number of images from 10 seedlings for each genotype and treatment. N in (i)=6 biologically independent samples. Experiments were repeated at least three times with similar results. Significance in (d‐h) was assessed by two‐way ANOVA with Tukey's multiple comparisons test, and in (i) by one‐way ANOVA with Tukey's multiple comparisons test. Different letters indicate significant differences at P<0.05.

Next, we used various biochemical approaches to investigate whether the MDP25/VDAC3 interaction directly affects MDP25 function. A high‐speed cosedimentation assay was conducted to assess the actin‐binding activity of MDP25. As shown in Figure S13b, the amount of MDP25 cosedimented with actin was similar in the presence or absence of VDAC3, suggesting that VDAC3 does not affect MDP25's ability to bind to actin filaments (Figure S13b). Furthermore, we used total internal reflection fluorescence (TIRF) microscopy to visualize actin dynamics in real time. Prepolymerized Oregon‐green‐labeled actin filaments adhered to the cover glass of a perfusion chamber [17]. MDP25 proteins were perfused into the chamber, and time‐lapse images were captured. Actin alone or in the presence of different VDAC3 variants showed minimal breakage (Figure S13c–e,i,j; Movie S10). The addition of MDP25 induced breaks along the filaments, demonstrating severing activity (Figure S13f, i,j; Movie S11). When VDAC3 was added to the reaction, the severing activity of MDP25 was reduced. However, the impacts of VDAC3 on MDP25 was abolished by its D168 mutation (Figure S13g–j; Movie S11). These data suggested that MDP25's function in regulating actin dynamics can be modified by interacting with VDAC3. Based on these biochemical results, one might predict opposite phenotypes between mdp25 and vdac3‐1 mutants if VDAC3 functions solely as a negative regulator of MDP25's actin‐severing activity; for example, vdac3‐1 mutants would be expected to mimic MDP25 overexpression. Instead, the genetic phenocopy of vdac3‐1 and mdp25 mutants suggests that the predominant in vivo role of VDAC3 is to ensure the proper positioning and functional integration of MDP25 at mitochondria‐actin contact sites, rather than acting as a global inhibitor. In the absence of VDAC3, MDP25 becomes mislocalized and is unable to support localized actin remodeling, resulting in a loss‐of‐function phenotype rather than simple hyper‐severing.

The functional significance of VDAC3 D168 was further assessed in vivo by expressing VDAC3 D168G in the vdac3‐1 mutant background. Under resting conditions, expressing VDAC3 D168G complemented the larger mitochondria phenotypes in the vdac3‐1 mutant as expressing WT VDAC3 did (Figure S4e). However, flg22‐induced changes in mitochondrial morphology and dynamics were not restored to WT level in VDAC3 D168G/vdac3‐1 as in VDAC3/vdac3‐1 (Figure 6d–f; Figure S4e). We also analyzed the actin organization in WT, vdac3‐1 mutants, and different VDAC3 complementation lines. Expression either WT VDAC3 or VDAC3 D168G can restore the actin defects in vdac3‐1 mutants under mock conditions. However, the actin responses to flg22 remained compromised in VDAC3 D168G/vdac3‐1 background (Figure 6g,h; Figure S14). Moreover, VDAC3 D168G expression failed to restore the compromised disease resistance in vdac3‐1 mutants as WT VDAC3 (Figure 6i). Collectively, these results provided evidence that MDP25/VDAC3 module is required for actin and mitochondrial responses following flg22 treatments.

3. Discussion

In this study, we investigated the role of host actin reorganization in mitochondrial remodeling during plant innate immunity. Flg22 treatment induces increased mitochondrial fusion activity, resulting in the excessive mitochondrial elongation. These dynamic changes in mitochondrial morphology facilitate the mixing of mitochondrial matrix components, which may promote the functional recovery of damaged mitochondria, as well as mROS and ATP production under biotic stresses. By monitoring the spatiotemporal dynamics between the actin cytoskeleton and mitochondria in real time, we show that mitochondrial elongation requires the assembly of actin bundles induced by flg22. We speculate that these actin bundles likely provide structural support for mitochondrial fusion and may help prevent already elongated mitochondria from undergoing fission. We further dissected the role of MDP25/VDAC3 complex in actin‐mediated mitochondrial elongation. Upon flg22 stimulation, MDP25 translocates from the PM to the cytosol. VDAC3 recruits cytosolic MDP25 and anchors it at peri‐mitochondrial actin contact sites. We hypothesize that at these sites, the MDP25/VDAC3 module cooperatively organizes actin remodeling to support mitochondrial fusion and elongation (Figure 7). Together, our findings provide insights into the molecular mechanism and biological significance of actin‐dependent mitochondrial dynamics during plant innate immune responses.

FIGURE 7.

A model depicting key findings in this work. Upon flg22 treatment, MDP25 translocates from the PM to the cytosol. VDAC3 recruits cytosolic MDP25 and anchors it at peri‐mitochondrial actin contact sites, where the MDP25/VDAC3 module cooperatively organizes actin remodeling to support mitochondrial fusion and elongation. This mitochondrial remodeling facilitates the exchange of mitochondrial contents for the functional recovery of damaged mitochondria and promotes ATP and mitochondrial ROS production for effective immunity in plant cells.

Mitochondrial dynamics play an important role in innate immunity in animals [52]. For instance, mitochondrial elongation is associated with enhanced antiviral immunity, while mitochondrial fragmentation has the opposite effect [53, 54, 55]. Although evidence in plants is limited, elongated mitochondria have been linked to enhanced resistance to fungal pathogens in rice [11]. In this study, we provide evidence that mitochondrial elongation is beneficial for plant defense against bacterial infection. During the initial 2 h of flg22 treatment, we found that mitochondria exhibit temporary damage, indicated by a sudden decrease in mitochondrial membrane potential. The membrane potential gradually recovers from 2 h onward, correlating with mitochondrial elongation. We hypothesize that active fusion‐induced mitochondrial elongation allows normal mitochondria to compensate for damaged ones. Through fusion, essential functional and structural components, such as proteins and lipids, can be diffused and shared between the two types of mitochondria, repairing dysfunction and mitigating environmental stress effects on mitochondria [56, 57]. However, the significant rescue of disease resistance by MIDI treatment in the mdp25 and vdac3 mutants suggests that mitochondrial elongation also contributes to immunity independently of fusion‐mediated content exchange. Although the precise contribution of elongated mitochondria to immunity remains unclear, we speculate that elongated morphology provides functional benefits such as stabilizing membrane potential, supporting sustained ATP production, and preventing excessive fragmentation into dysfunctional units [58]. Therefore, it is likely that both fusion‐dependent content exchange and elongation‐dependent functional stabilization contribution to immune responses. The importance of mROS in plant immunity has been long recognized. Our study and others reported enhanced mROS production upon flg22 treatment, and depletion of mROS pharmaceutically or genetically can compromise plant disease resistance [2, 9]. However, how mitochondrial elongation mechanistically enhances mROS and ATP production remains an important open question.

We observed that mitochondrial elongation transiently occurs from 6 to 12 h following flg22 treatment. This temporal pattern likely reflects a transient, stress‐adaptive response triggered by MAMP perception. It is well established in animal cells that mitochondrial function and dynamics are highly sensitive to immune activation [59]. Although less is known in plant cells, we propose that 6‐12 h may represent a critical phase during PTI, in which mitochondria are highly activated to support energy and redox demands of defense responses. At earlier stages (e.g., 1–6 h), immune signaling may not have fully propagated to mitochondria, and the corresponding metabolic and redox adjustments may still be in progress. Between 6 and 12 h, flg22‐triggered signaling promotes mitochondrial fusion to support mitochondrial integrity and function needed for defense, as reflected by the observed increase in mitochondrial membrane potential, mROS, and ATP levels (Figure 3). By 12 h, it is possible that homeostatic feedback mechanisms may begin to downregulate mitochondrial activity to prevent excessive oxidative stress and energy imbalance. Sustained signaling may also trigger mitochondrial fission or mitophagy pathways, thereby reversing the elongation phenotype. Further investigation will be required to test these possibilities.

We showed that flg22‐induced mitochondrial elongation is compromised in fls2 mutant. Given that flg22 perception by FLS2 is the initial step for activating downstream signaling [60], it is likely a prerequisite for activating MDP25/VDAC3‐dependent mitochondrial remodeling. Although the specific signals downstream of FLS2 activation ultimately engage the MDP25/VDAC3 machinery to drive fusion remains unresolved in this study, our data showed that MDP25 translocation from the PM to the cytosol requires Ca²⁺ signaling, suggesting that flg22‐induced Ca²⁺ transients are likely an important link between FLS2 activation and MDP25/VDAC3‐mediated mitochondrial fusion. It is also likely that other PTI signals may contribute to engaging this machinery, and future studies will be needed to define the precise molecular relationships.

It was previously reported that loss of VDAC1 or VDAC4 in Arabidopsis results in swollen mitochondria [33]. Consistent with this, our results showed that mutation of VDAC3 also leads to an increase in mitochondrial sizes (Figure S4d). Although the function of plant VDACs remain poorly investigated, it is likely that VDAC depletion results in mitochondrial swelling via a buildup of mitochondrial metabolites, consequently leading to impaired membrane potential and ATP synthesis. Unlike VDAC3 mutation, reduced MDP25 levels in cells can compromise mitochondrial activity without affecting their morphology. Previous studies have demonstrated that disruption of actin dynamics by treatments with LatB or jasplakinolide, as well as the loss of actin‐nucleating factor Arp2/3 complex, leads to a decrease in mitochondrial membrane potential [25, 26]. Therefore, the impaired actin dynamics likely contribute to the defects in mitochondrial activity observed in the mdp25 mutant. Although this study highlights the role of MDP25 and VDAC3 in mitochondrial function and dynamics during defense responses, it is likely that these proteins regulate plant immunity through additional, yet‐to‐be‐identified mechanisms. Nevertheless, we found that pharmacological induction of mitochondrial elongation in mdp25 and vdac3 mutants partially rescues their compromised disease resistance. This finding suggests that MDP25 and VDAC3 contribute to plant immunity, at least in part, by promoting mitochondrial elongation.

Our finding that the VDAC3^D168G mutant, which disrupts the VDAC3‐MDP25 association, can restore the actin defects of vdac3 under resting conditions but not the flg22‐induced actin response (Figure 6g) suggests that VDAC3 regulates actin organization through mechanisms beyond its interaction with MDP25. Although VDAC3 is localized to the mitochondrial outer membrane and has not been reported to directly modulate actin, accumulating evidence indicates that mitochondria communicate both functionally and physically with the actin cytoskeleton [19]. Thus, VDAC3 may influence actin dynamics indirectly, for example by interacting with other actin regulators or by affecting mitochondrial functions such as ATP production or Ca²⁺ homeostasis. Consistent with this view, impaired mitochondrial activity has been shown to cause defects in actin dynamics [61]. Together, these findings highlight the complex regulatory interplay between mitochondria and the actin cytoskeleton.

The precise relationship between MDP25‐mediated actin severing and actin bundle formation remains unclear. However, our study and others have consistently shown that altered severing activity is often associated with changes in actin bundle formation [62, 63, 64]. We propose that the effect of MDP25 on actin bundling is likely indirect. Actin severing generates new filament ends that can serve as nucleation sites for subsequent filament elongation. Therefore, the reduced severing activity in the mdp25 mutant would limit the production of free barbed ends, ultimately reducing the number of newly formed filaments available for bundling.

The identification of multiple VDAC isoforms as MDP25‐associated proteins raises the possibility that VDAC family members may contribute redundantly to immune regulation. Although our current data support a primary role for VDAC3, the shared structural features and overlapping mitochondrial localization of VDAC isoforms suggest that additional members of the family may also participate in immune signaling [34]. The extent of such functional redundancy remains an open question for future investigation.

4. Methods

4.1. Plant Materials and Constructs

All experiments were conducted using the Arabidopsis thaliana Columbia ecotype (Col‐0). Surface‐sterilized seeds were plated onto ½ Murashige and Skoog (MS) medium supplemented with 1% sucrose and stratified at 4°C for 2 d. Plants were grown in a growth chamber under long‐day conditions (16 h light/8 h dark) at 22°C.

The mdp25 mutant and various MDP25‐GFP transgenic lines were generated in previous studies [28, 29]. The T‐DNA mutants of vdac3‐1 (SALK_127899) were obtained from the Nottingham Arabidopsis Stock Centre. To generate VDAC3 or VDAC3 D168G overexpression lines, the full‐length WT or mutated cDNA of VDAC3 was amplified and cloned into the binary vector pCAMBIA1390 containing GFP or mCherry, the resulting constructs were introduced into vdac3‐1 mutant background by Agrobacterium‐mediated transformation. To visualize mitochondria, the mitochondrial targeting sequence, the first 29 aa of yeast (Saccharomyces cerevisiae) cytochrome c oxidase IV (ScCOX4), was fused to the N‐terminus of Dendra2, GFP, or mScarlet, and cloned into the binary vector pCAMBIA1390. The corresponding constructs were introduced into WT background to generate stable transgenic line expressing mt‐Dendra2, mt‐GFP and mt‐mScarlet. Transformants were selected by antibiotic resistance and fluorescence intensity. VDAC3‐mCherry complementation lines were crossed with MDP25‐GFP complementation lines to generate dual‐color plants expressing both MDP25‐GFP and VDAC3‐mCherry. To simultaneously visualize mitochondria and actin cytoskeleton, mt‐mScarlet was crossed to GFP‐fABD2 line, and the F2 plants were used for imaging. The mdp25 and vdac3‐1 mutants were crossed to WT plants expressing both mt‐mScarlet and GFP‐fABD2 to generate dual marker line in the mutant background. Homozygous mutants, as well as WT siblings, were recovered from F2 populations. The mCherry‐fABD2 or GFP‐fABD2 reporter was introduced into the mutant and complementation lines by crossing. To generate constructs for the pull‐down assay, the full‐length WT or mutated cDNA of VDAC3 was cloned into pET28a to generate His‐tagged VDAC3 or VDAC3 D168G. The MDP25 cDNA was cloned into pGEX‐6p‐1 to generate an N‐terminal GST fusion. For the split‐luciferase complementation assay, the WT or mutated cDNAs of VDAC3 and MDP25 were cloned into Cluc‐pCAMBIA1300 or Nluc‐pCAMBIA1300, respectively. All primers used in this study are listed in Table S3.

4.2. Chemical Treatments

Chemicals used in this study include LatB (Sigma‐Aldrich), oryzalin (Sigma‐Aldrich), flg22 (Sangon Biotech, Shanghai), elf18 (Sangon Biotech, Shanghai), chitin (Sigma‐Aldrich), flgII‐28 (Sangon Biotech, Shanghai), MitoSOX Red (Invitrogen), MitoTracker (Invitrogen), TMRM (Sigma‐Aldrich), FM4‐64 (Invitrogen), MIDI and LaCl₃ (Sigma‐Aldrich). LatB, MitoSOX Red, MitoTracker, TMRM, MIDI, oryzalin, and FM4‐64 are dissolved in DMSO, while flg22, elf18, flg‐II28, chitin, and LaCl₃ are dissolved in distilled water. For live‐cell imaging analysis, 2‐week‐old seedlings were spray‐treated with 10 µM flg22, 10 µM elf18, 10 µM flgII‐28, or 500 µg/mL chitin. After 5‐min incubation, the excess solution was then removed, and seedlings were transferred to ½ MS agar plates for 12 h or the indicated time periods before imaging. LatB treatments were performed at 5 µM for 30 min. Oryzalin treatments were performed at 300 µM for 15 min. For MIDI treatment, 2‐week‐old seedlings were transferred to ½ MS agar plates supplemented with 2 µM MIDI for 6 h prior to imaging or bacterial infection. The concentration and duration of MIDI treatment were chosen based on our preliminary time‐course observations and following the practical guidelines described in Yang et al. (2023) [44], both of which indicated that this treatment is sufficient to induce robust mitochondrial elongation without causing obvious cytotoxicity. For mitochondrial labeling, 2‐week‐old seedlings were stained with 500 nM MitoTracker for 1 h. To analyze mitochondrial membrane potential, seedlings were incubated with 500 nM TMRM for 30 min at room temperature. To analyze mROS, MitoSOX Red was applied to cotyledons of 2‐week‐old seedlings expressing mt‐GFP at 10 µM for 1 h.

4.3. Co‐IP Assay

Full‐length cDNA of VDAC3 was cloned into the GFP tag and MDP25 was cloned into the mCherry tag to generate U10:VDAC3‐GFP and U10:MDP25‐mCherry. These resulting constructs were infiltrated into N. benthamiana via the Agrobacterium‐mediated method. After 48 h, total proteins were extracted for coIP using extraction buffer and incubated with anti‐GFP agarose beads for 2 h. Beads were washed eight times with washing buffer (50 mM HEPES, pH 7.5, 150 mM KCl, 1 mM EDTA, 0.5% Triton X‐100, 1 mM DTT). After washing, the immunoprecipitated proteins were separated by SDS‐PAGE and detected by anti‐GFP and anti‐mCherry immunoblotting.

4.4. FRET‐FLIM Assay

Leaves from 7‐d‐old seedings expressing both VDAC3‐GFP and MDP25‐mCherry were used 3 h after treatment with mock, 10 µM flg22 in the presence or absence of 10 mM LaCl₃. FRET‐FLIM analysis was performed using an inverted FV1200 confocal fluorescence microscope (Olympus) equipped with an oil immersion objective (60×, numerical aperture = 1.3). The donor fluorophore (GFP) was excited with 480 nm laser and detected with 520/35 nm bandpass filter. In all experiments, the laser power was adjusted to achieve average photon counting rates ≤105 photons per second. Data acquisition and analysis were performed with SymphoTime 64 software (PicoQuant).

4.5. Cytosolic and PM Fractionation and Immunoblot Analysis

Seven‐day‐old MDP25‐GFP seedlings were treated for 3 h with either mock, 10 µM flg22, or 10 µM flg22 combined with 10 mM LaCl₃. Following treatment, seedlings were harvested for protein extraction to assess the subcellular localization of MDP25. PM proteins were isolated using the Minute Plasma Membrane Protein Isolation Kit for Plants (Invent Biotechnologies), according to the manufacturer's instructions. MDP25‐GFP levels in the cytosolic and PM fractions were detected via immunoblotting using an anti‐GFP antibody. PM H⁺‐ATPase and Rubisco were used as markers for PM and cytosolic fractions, respectively. H⁺‐ATPase was detected with an anti‐H⁺‐ATPase antibody (Agrisera).

4.6. GST Pull‐Down Assay

GST or GST‐MDP25 were expressed in E. coli and purified using the glutathione agarose beads (Cytiva). His‐tagged VDAC3 or VDAC3 D168G fusion proteins were extracted and purified with Ni‐NTA His bind resin (Cytiva). For the GST pull‐down assay, 5 µg GST or GST‐MDP25 and 5 µg His‐VDAC3 or His‐VDAC3 D168G were incubated with 30 µL glutathione agarose beads in binding buffer (140 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO4, 1.8 mM KH₂PO4, 0.5% NP‐40) for 1 h. The beads were washed seven times with washing buffer (40 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO4, 1.8 mM KH₂PO4, 0.5% NP‐40). The bound protein was eluted with 10 mM glutathione, separated by SDS‐PAGE, and detected by immunoblotting with antibodies against His or GST.

4.7. Split‐luciferase Complementation Assay

Agrobacterium tumefaciens GV3101 containing the VDAC‐Cluc and MDP25‐Nluc plasmids were infiltrated into expanded leaves of N. benthamiana and incubated in the growth room for 48 h before the LUC activity measurement. For CCD imaging and LUC activity measurement, 1 mM luciferin was sprayed onto the leaves. A cooled CCD imaging apparatus was used to capture the LUC image.

4.8. GFP Pull‐Down Assay and Mass Spectrometry

A procedure described in a previous study was adopted to identify MDP25‐interacting proteins [65]. Briefly, we collected 1.5 g of transgenic plants expressing GFP or the mdp25 mutant expressing pMDP25: MDP25‐GFP at 10 d old. The plant samples were rapidly frozen and ground in liquid nitrogen. Total protein was extracted using the IP buffer (50 mM HEPES, pH 7.5, 150 mM KCl, 1 mM EDTA, 0.5% Triton X‐100, 1 mM DTT, proteinase inhibitor cocktail). Debris was removed from the lysate by centrifugation at 16,000 g for 15 min. The supernatant was filtered through a 0.22 mm low‐protein binding filter (Millipore) and incubated with 50 µL anti‐GFP agarose beads for 2 h. Beads were collected at 500 g for 5 min. The sample proteins were loaded onto a single lane on an SDS‐PAGE gel after washing the GFP beads five times. The quality of the immunoprecipitates was assessed by Coomassie brilliant blue staining and western blotting using anti‐GFP. The immunoprecipitates were separated using SDS‐PAGE, and digested into peptides to identify MDP25 interactors by LC‐MS/MS analyses.

4.9. FRET Imaging and Analysis of ATeam 1.03‐nD/nA

Fluorescent images were collected from cotyledon pavement cells from 2‐week‐old seedlings expressing ATeam 1.03‐nD/nA using a LSM710 confocal microscope (Carl Zeiss) equipped with a 63x 1.4NA oil‐immersion lens. Ateam1.03‐nD/nA fluorescence was excited at 458 nm, and the emission signals were detected at 470–507 nm (Em463‐507, CFP image) and 526–561 nm (Em524‐544, YFP image), respectively. Fluorescence intensity was measured using ImageJ, and the YFP/CFP ratio was calculated as the FRET efficiency.

4.10. TEM

To analyze the mitochondrial morphology, detached cotyledons of 2‐week‐old seedlings were fixed with 2% paraformaldehyde (PFA), 2.5% (w/v) glutaraldehyde (Sigma‐Aldrich), 0.1 M phosphate buffer (50 mM Na₂HPO₄, 50 mM NaH₂PO₄, pH 7.4) for 1 h at room temperature, then held overnight at 4°C. The samples were washed with 0.1 M phosphate buffer for 20 min three times and post‐fixed in 1% (w/v) OsO4 (Ted Pella) and 0.8% (w/v) potassium ferrocyanide (Sigma‐Aldrich) for 2 h. After washing with distilled water for three times, samples were dehydrated through a graded alcohol series and embedded in Embed 812 resin (EMS). Resin blocks were cut into 70‐nm sections using an ultramicrotome (Leica Microsystem, UC7) for TEM (JEOL, JEM‐1400).

4.11. Live‐Cell Imaging

Live‐cell imaging was conducted on cotyledon pavement cells from 2‐week‐old seedings after indicated treatment. Detached cotyledons were mounted in the desired solutions between a slide and a coverslip on a chambered slide, which was made from two parallel strips of ultrathin double‐sided adhesive tape for imaging. Z‐stacks from the surface to the mid‐plane were collected to analyze mitochondrial volume and density. For analysis of mitochondrial length‐to‐width ratios or dynamics, single optical sections were collected at the cell surface. To minimize selection bias, multiple fields of view were randomly scanned for each sample rather than any subjective assessment of signal quality. Laser power, gain, and exposure times were kept consistent across all samples within each experiment to ensure comparability. For each genotype or treatment, multiple independent biological replicates were imaged, and multiple cells were quantified per replicate to ensure robustness. Image collection, analysis and quantification were carried out in a double‐blinded manner, with the experimenter unaware of the genotypes or treatments, to ensure objectivity and minimize bias. Time‐lapse images were taken on an airyscan LSM880 confocal microscope (Carl Zeiss) equipped with 63x 1.4NA oil‐immersion lens at 0.21‐s intervals for 21 s or 105 s. To determine actin array organization, image stacks with a z‐step of 0.5 µm were captured using the LSM880 confocal microscope (Carl Zeiss) equipped with 63x 1.4NA oil‐immersion lens. For the analysis of single actin filament dynamics, images were taken on ELYRA 7 TIRF microscope (Carl Zeiss) equipped with 100x 1.46NA oil‐immersion lens at 0.2‐s intervals for 100 s. To analyze the dynamics of both actin and mitochondria, two‐color images were collected on the spinning disk confocal microscope (UltraView VoX, PerkinElmer), equipped with a CSU‐X1 Yokogawa spinning disk head fitted to a Nikon TiE inverted microscope, a Hamamatsu EMCCD 9100‐13, and a 100x 1.49NA oil‐immersion objective. Single optical sections were taken at cell cortical regions at 1.36‐s intervals for 5 min. Excitation and emission wavelengths for the different probes were as follows: GFP, 488/500 to 550 nm; MitoSOX Red, 510 to 580‐610 nm; mCherry/mScarlet/TMRM/FM4‐64, 561/570 to 620 nm. Image acquisition was controlled by ZEN software (Zeiss) or Volocity Demo 6.1.1 (PerkinElmer).

4.12. Quantitative Analysis of Actin Dynamics

The percentage of occupancy (density) were quantified as previously described [13]. To quantify the relative fluorescent intensity level of actin bundles in the cortical array, the total fluorescence pixel intensity of each filamentous structure, after subtraction of the background intensity, was measured in a cell and was further normalized to the intensity of single actin filaments. The faintest actin structures in the cell were hand‐selected and assumed to be single actin filaments. The severing frequency was quantified as previously described [66]. For nucleation and bundling frequency analysis, a 125 and 49‐µm² ROI was randomly selected, respectively. All observable events were counted during a 100‐s time period [67].

4.13. Quantification of Mitochondrial Morphologies and Dynamics

Live‐cell images were collected from cotyledon pavement cells of 2‐week‐old seedlings expressing mt‐GFP or stained with Mitotracker after treatment with mock or flg22 for the indicated time periods. To quantify the ratio of length and width of mitochondria, the length and width of each mitochondrion in each image were measured in ImageJ. To quantify the mitochondrial density, z stacks were captured. Cells were outlined by FM4‐64 staining and the number of mitochondria in each cell was manually counted. To quantify the mitochondrial volume, 2‐week‐old seedlings expressing mt‐GFP were treated with mock or 10 µM flg22 for 12 h. Cotyledons were then detached and fixed in 2% paraformaldehyde for 1 min. Image stacks with a z‐step of 0.5 µm were collected and projected with maximum intensity, and the volume of each mitochondrion was measured using the 3D Objects Counter plugin in ImageJ. Analyses of mitochondrial dynamics were performed on time‐lapse images. The behaviors and trajectories of mitochondria were manually tracked.

To analyze actin and mitochondria dynamics, two‐color imaging was performed on cotyledon pavement cells of 2‐week‐old seedlings expressing mt‐mScarlet and GFP‐fABD2. To determine the percentage of fusion events occurring either on or off actin bundles, we manually counted the fusion events that led to the formation of mitochondria of specified length in a continuous 5‐min image sequence. The count of mitochondria on actin bundles was conducted within a 90‐µm² ROI. Mitochondria stay on actin bundles for more than 3 consecutive frames were selected to quantify the association frequency and duration of mitochondria on actin bundles. For the quantification of mitochondria‐adjacent actin dynamics, mitochondria that remained associated with actin filaments for at least five consecutive frames were selected for analysis. For each mitochondrion, a 100‐µm² ROI surrounding the mitochondrion was defined. Within this ROI, severing events of all actin filaments were tracked over time, from the start of observation until the filaments were completely severed. The severing frequency was defined as the number of these events per unit area per unit time (events µm⁻² s⁻¹).

Exchange of matrix‐localized mt‐Dendra2 [68, 69] was analyzed between mitochondria in cotyledons of 2‐week‐old seedlings. Single optical slices were collected with an airyscan LSM880 confocal microscope (Carl Zeiss) equipped with 63x 1.4NA oil‐immersion objectives. The 488 nm laser and the 561 nm laser excited Dendra2 in the unconverted state and photo‐converted state, respectively. Photoconversion was performed at 1.82 fps (pixel dwell time of 0.8 µs) (or a scan speed of 0.8 µs/pixel) within a rectangular ROI using the 405‐nm laser at 4% power output and for a total of 8.25 s (90 bleaching iterations). After photoconversion, image capture continued for 1182 loops equating to 10.84 min. Green and red channels were captured simultaneously preconversion and postconversion using a 488‐ nm laser and a 561‐nm laser, respectively.

4.14. High‐Speed Cosedimentation Assay

Actin was dialyzed overnight against buffer G (5 mM Tris‐HCl, 0.1 mM CaCl₂, 1 mM NaN₃, 0.2 mM ATP·2Na, 0.5 mM DTT, pH 7.0). Before use, MDP25, VDAC3, and actin were clarified by centrifugation at 55 000 g for 1 h. Actin was prepolymerized in 1× KMEI buffer (50 mM KCl, 1 mM MgCl₂, 1 mM EGTA, 10 mM imidazole, pH 7.0) at room temperature for 2 h. MDP25 and VDAC3 was then incubated with preformed actin filaments at a 1:3 ratio in a 100‐µL reaction volume. Following incubation, the reaction mix was centrifuged for 1 h at 55 000 g. The supernatant (80 µL) was transferred to a separate tube, and 16 µL of 6× protein loading buffer was added to it. The pellet was suspended by 100 µL buffer G, and 20 µL of 6 × protein loading buffer was added. The samples were then separated by 12% SDS‐PAGE and stained with Coomassie Brilliant Blue R (Sigma‐Aldrich).

4.15. TIRF Microscopy

Individual severing events along actin filaments were imaged by time‐lapse TIRF microscopy. The assembly of monomeric actin (Oregon‐green labeled) was initiated by the addition of one‐tenth volume of 10 × KMEI (0.5 mM KCl, 10 mM MgCl₂, 10 mM EGTA, 0.1 mM Imidazole, pH 7.0). Actin filaments (15%–50% Oregon‐green‐actin, 0.125–1 mM) were mixed with 2× TIRF buffer (20 mM imidazole [pH 7.4], 100 mM KCl, 2 mM MgCl₂, 2 mM EGTA, 0.4 mM ATP, 10 mM DTT, 30 mM glucose, 40 mg ml⁻¹ catalase, 200 µg ml⁻¹ glucose oxidase, and 1% methylcellulose) and transferred to a microscope flow chamber for imaging at room temperature. MDP25 and VDAC3 proteins were introduced after placing the chamber on the microscope stage. TIRF images were collected at 0.5 s intervals using an ELYRA 7 (Carl Zeiss). Microscope slides (24 × 50 mm 12‐545‐F; Fisher Scientific) and cover‐slips (22 × 22 mm 12‐542‐B; Fisher Scientific) were cleaned using piranha solution (a 3:1 mixture of sulfuric acid and 30% hydrogen peroxide) overnight. The glass was then incubated with 2 mg ml⁻¹ methoxy‐PEG‐silane MW 5000 and 2 mg ml⁻¹ biotin‐PEG‐silane MW 3400 (Laysan Bio) in 95% ethanol (pH 2.0) at 70°C. Two parallel strips of double‐sided tape were placed on both ends of the coverslip to create a flow chamber.

The stack of 780 frames from each time‐lapse series was used to observe and count breaks along single filaments in all images following the addition of protein. The maximum length of each filament was measured on the first frame and subsequent breaks recorded over time until the filament disappeared. Severing frequency for each actin filament was calculated as the number of breaks, per unit filament length, per unit time (i.e., breaks/µm/s). The severing frequency for a particular protein in a certain condition was estimated by calculating the average severing frequency from filaments that were longer than 10 µm. Three independent batches of each protein and three technical replications were used per condition.

4.16. PTI Response Analyses

For MAPK activation assay, 2‐week‐old seedlings were soaked in water containing flg22 and frozen in liquid nitrogen. Total protein was extracted using the extraction buffer (50 mM HEPES, pH 7.5, 150 mM KCl, 1 mM EDTA, 0.5% Triton X‐100, 1 mM DTT, proteinase inhibitor cocktail). Protein extracts were probed with anti‐MPK6 (Sigma‐Aldrich) to assess MPK6 protein levels, or anti‐p44/42 MAPK (Cell Signaling Technology) to determine the phosphorylation of MPK3, MPK4 and MPK6. For the oxidative burst assay, leaf disks (4‐mm diameter) from 4‐week‐old plants were floated on water overnight in a 96‐well plate (one disk per well) and then treated with 1 µM flg22 in 100 µL buffer containing 20 µM luminol and 10 µg ml⁻¹ horseradish peroxidase. The luminescence was recorded using a FlexStation3 (Molecular Devices). For callose deposition analyses, 2‐week‐old seedlings were incubated with 1 µM flg22 for 24 h. To stain callose, seedlings were incubated for at least 24 h in 95% to 100% ethanol until all tissues were transparent, washed in 0.07 M phosphate buffer (pH 9.0), and incubated for 1 to 2 h in 0.07 M phosphate buffer containing 0.01% aniline blue (Sigma‐Aldrich). Images of callose deposits from whole cotyledons were collected using a Zeiss ImagerM1 epifluorescence microscope equipped with a 20x 0.5NA PlanFluor objective under ultraviolet light. The number of callose spots was quantified using ImageJ. For stomatal aperture assay, 2‐week‐old seedings were incubated in stomata‐opening buffer (50 mM KCl, 10 µM CaCl₂, and 10 mM MES, pH 6.15) for 2 h in a growth chamber at 22°C under constant illumination. Then seedlings were transferred to stomata‐opening buffer supplemented with mock or 1 µM flg22. Stomatal apertures were measured after treatment with mock, 1 µM flg22 for 1 h using ImageJ.

4.17. Bacterial Growth Assay

To assess bacterial growth, 2‐week‐old seedlings were spray inoculated with DC3000 suspensions at a concentration of 1 × 10⁶ colony‐forming units per ml. At 48 h post‐infection, six seedlings were harvested for each genotype and ground in 10 mM MgCl₂. After extracting bacteria from the plant material, serial dilutions of leaf extracts were prepared. A 2‐µL aliquot from each dilution was plated to enumerate bacterial colonies. Each data point represents the average bacterial count from six replicates.

4.18. Statistical Analysis

Statistical analysis, including Student's t‐test, and one‐way or two‐way ANOVA with Tukey's multiple comparisons test was carried out using GraphPad Prism 9.5.1. Differences were considered significant when p values were <0.05. Data represent the mean ± SD. Details of statistical analysis are provided in the figure legends.

Author Contributions

J.H. and J.L. designed the experiments. J.H., P.L., X.C., L.L., and Q.P. performed experiments. J.H., P.L., X.C., L.L., Q. P., and J.L. analyzed the data; J.H. and J.L. wrote the paper. All authors commented and agreed on the manuscript before submission.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File 1: advs73563‐sup‐0001‐SuppMat.docx.

Supporting File 2: advs73563‐sup‐0002‐MovieS1.mp4.

Supporting File 3: advs73563‐sup‐0003‐MovieS2.mp4.

Supporting File 4: advs73563‐sup‐0004‐MovieS3.mp4.

Supporting File 5: advs73563‐sup‐0005‐MovieS4.mp4.

Supporting File 6: advs73563‐sup‐0006‐MovieS5.mp4.

Supporting File 7: advs73563‐sup‐0007‐MovieS6.mp4.

Supporting File 8: advs73563‐sup‐0008‐MovieS7.mp4.

Supporting File 9: advs73563‐sup‐0009‐MovieS8.mp4.

Supporting File 10: advs73563‐sup‐0010‐MovieS9.mp4.

Supporting File 11: advs73563‐sup‐0011‐MovieS10.mp4.

Supporting File 12: advs73563‐sup‐0012‐MovieS11.mp4.

Supporting File 13: advs73563‐sup‐0013‐Data.zip.

Data Availability Statement

The authors declare that all data supporting the findings of this study are available within the manuscript and within its supplemental materials. Source data are provided with this paper. Proteomic datasets can be accessed via the following link: https://www.iprox.cn/page/PSV023.html;?url=1737009076636PC0u. All raw confocal images for main figures can be accessed via the following link: https://data.mendeley.com/preview/hwry4k6zrt?a=8b70c13a‐37eb‐42a3‐b9e8‐d4d85cf6e034; all raw confocal images for supplemental figures can be accessed via the link: https://data.mendeley.com/preview/78w98grrdw?a=22959948‐8ad0‐41b5‐af57‐9521fa0869e8. The original, uncropped western blot and SDS‐PAGE images are provided in the source data file.