Regulation of Oomycete Autophagy, Lipid Droplet Accumulation and Pathogenesis by Three Rab GTPases

ABSTRACT

Among eukaryotes, Rab GTPases are critical for intracellular membrane trafficking and possess various functions. Oomycetes, responsible for many devastating plant diseases, pose a significant threat to global agriculture. However, the functions of Rab GTPases in oomycetes are largely uncharted. In this study, we functionally characterised two Rab8 homologues, PlRab8A and PlRab8B, from a plant‐pathogenic oomycete, Peronophythora litchii. These genes are crucial for mycelial growth and play significant roles in the production and morphology of sporangia, as well as the formation of zoospores. PlRab8A and PlRab8B also contribute to maintaining cell wall integrity, osmotic stress tolerance and oospore formation, which are vital for the organism's survival and virulence. Moreover, PlRab8A and PlRab8B are essential for the full virulence of P. litchii by influencing laccase activity. We also found that PlRab8B co‐localised with lipid droplets (LDs) and negatively affected LD accumulation. Autophagy was enhanced and autophagy‐related genes (PlATGs) were up‐regulated in PlRab8B mutants. Additionally, PlRab8A was found to interact with PlRab6. Similar to PlRab8, PlRab6 is also required for growth, zoospore formation and pathogenicity of P. litchii. Taken together, this study reported for the first time the functions of three Rab proteins regulating autophagy, LD accumulation and pathogenesis in oomycetes.

PlRab6– PlRab8A complex controls growth, virulence and autophagy, while PlRab8B modulates sporulation and virulence via autophagy and LD metabolism in P. litchii.

1. Introduction

Oomycetes cause devastating agricultural diseases, including potato late blight ( Phytophthora infestans ), soybean root rot (Phytophthora sojae) and litchi downy blight (Peronophythora litchii) (Punia and Kumar 2021; Hu et al. 2022). Among these diseases, litchi downy blight is notable for causing significant yield losses, with an annual reduction of 30% to 80% (Situ et al. 2022). The pathogen's rapid spread through zoospores and oospores is particularly accelerated by temperature fluctuations that follow rainfall events (Xu et al. 2022). Although some basic biological characteristics of this important pathogen have been described, the molecular mechanisms associated with its development and pathogenicity remain largely unknown (Jiang et al. 2017; Situ et al. 2022). To further elucidate the pathogenic mechanisms of P. litchii, researchers have increasingly focused on fundamental cellular processes, particularly membrane trafficking, which is regulated by Rab GTPases—key molecular switches in vesicle transport and pathogen–host interactions (Situ et al. 2022). This process is essential in eukaryotic cells, as it not only affects their normal functioning but also plays a significant role in the infection and pathogenicity of pathogens.

Membrane trafficking, a fundamental eukaryotic process, governs vesicle transport between organelles and is critical for pathogen‐host interactions (Jenkins et al. 2020). Central to this process are Rab GTPases, molecular switches that cycle between GTP‐bound (active) and GDP‐bound (inactive) states to regulate vesicle budding, motility and fusion (Alexandrov et al. 1994; Ullrich et al. 1994; Grosshans et al. 2006; Stenmark 2009). Spatiotemporal regulation of vesicle traffic is ensured by cross‐talk between multiple Rab GTPases via shared effectors or recruitment of selective Rab activators (Stenmark 2009).

Rab proteins, which are ubiquitous in eukaryotic cells, play crucial roles in regulating development, protein secretion and pathogenicity, thereby attracting considerable research attention. In fungal pathogens, Rab GTPases are known to govern virulence‐related processes, including sporulation, effector secretion and host invasion (Powers‐Fletcher et al. 2013; Qi et al. 2014; Yang et al. 2017; Yan et al. 2020). For instance, mutants of Rab genes in Magnaporthe oryzae display defective conidiation and reduced pathogenicity (Qi et al. 2014; Yang et al. 2017). Despite these findings, the involvement of Rab GTPases in autophagy and interspecies crosstalk in oomycetes remains unexplored, which restricts the potential for targeting these pathways in disease control strategies (Yang et al. 2018; Yan et al. 2020). Additionally, while Rab proteins have been implicated in plant infection in M. oryzae and Fusarium graminearum and are well documented in model eukaryotes such as Saccharomyces cerevisiae (e.g., Ypt1 and Sec4/Rab8) (Tian et al. 2014; Liu et al. 2015; Zheng et al. 2015), their specific functions in pathogenic oomycetes, including their roles in autophagy and interspecies crosstalk, remain largely unknown.

The objectives of this study were to investigate the role of Rab GTPases in the development and pathogenicity of P. litchii. We found that Rab GTPases contributed to growth, stress tolerance, sporulation and pathogenicity in this organism. Additionally, PlRab8A interacted with PlRab6, while PlRab8B was involved in autophagy processes. This study provided evidence of crosstalk between Rab GTPases and autophagy in P. litchii, demonstrating their roles in growth, sexual and asexual development, as well as pathogenicity.

2. Results

2.1. Phylogenetic and Transcriptional Profile of Rab Genes in P. litchii

Rab proteins are known to play crucial roles in the morphological development of eukaryotic organisms (Zheng et al. 2015). However, the functions of oomycete Rab proteins are still unclear. We identified 36 Rab GTPase proteins in P. litchii with diverse transcriptional profiles. The heatmap analyses revealed that PlRab8A (Plitchii004269.m1) and PlRab8B (Plitchii008012.m1) possessed the typical RAB domain and displayed high expression levels during zoospore and infection stages (Figure S1). These features implied that they have crucial roles and make them prime candidates for functional investigation. Sequences were cloned from cDNA of P. litchii, and the coding sequence (CDS) regions of PlRab8A and PlRab8B were 609 and 624 bp in length, respectively.

To understand the evolutionary relationship of Rab8 among oomycetes, fungi, plants and animals, homologous proteins of PlRab8 were identified from Phytophthora fragariae, Ph. sojae, Ph. capsici, Ph. infestans, Ph. ramorum, Ph. cinnamomi, Ph. parasitica, Ph. palmivora, Pythium oligandrum, Py. brassicum, Desmarestia dudresnayi, Homo sapiens , Arabidopsis thaliana , Danio rerio and S. cerevisiae . Phylogenetic analysis revealed that oomycete Rab8 GTPases diverged into two major clades. PlRab8A and PlRab8B were placed into distinct clades (Figure 1a), both showing closer evolutionary relationships to animal ( H. sapiens , D. rerio ) and plant ( A. thaliana ) homologues than to fungi ( S. cerevisiae ). Notably, PlRab8A exhibited greater similarity to A. thaliana and algal homology, D. dudresnayi, than to PlRab8B. PlRab6 was evolutionarily distant from both PlRab8 isoforms, suggesting potential functional and evolutionary divergence among these three GTPases.

FIGURE 1.

The phylogenetic analysis, transcriptional profile and knockout of PlRab8A and PlRab8B. (a) Neighbour‐joining analysis using MEGA 11 with 1000 bootstrap replicates shown as percentages along the branches for PlRab8 homologues from various species (Ph. fragariae, Ph. sojae, Ph. capsici, Ph. infestans, Ph. ramorum, Ph. cinnamomi, Ph. parasitica, Ph. palmivora, Py. oligandrum, Py. brassicum, D. dudresnayi, H. sapiens, A. thaliana, D. rerio and S. cerevisiae ) (where Ph. = Phytophthora and Py. = Pythium). (b and c) Analysis of transcription levels for PlRab8A (b) and PlRab8B (c) during different stages of Peronophythora litchii life cycle including mycelium (MY), sporangia (SP), zoospore (ZO), cyst (CY), cyst germination (CG) and 1.5 to 24 h post‐infection (IF1.5–24 h). The P. litchii actin gene was used as internal control, and results were calculated using the 2^−ΔΔCt method. Data are mean ± SD based on three independent replicate experiments. Asterisks represent significant differences (t test, **p < 0.01; ***p < 0.001). (d) CRISPR/Cas9‐mediated knockout of the PlRab8 gene. The SgRNA1 and SgRNA2 were inserted into the pYF2.3G‐Ribo‐sgRNA vector, while the left and right arms of the PlRab8A or PlRab8B gene were inserted into the pBSSK vector. (e) PCR and electrophoresis analysis of Δplrab8a (T39, T50 and T54) and Δplrab8b (T12, T16 and T33). These experiments were independently repeated three times.

We analysed the transcripts of PlRab8A and PlRab8B at various life cycle stages, including mycelia (MY), sporangia (SP), zoospores (ZO), cysts (CY), cyst germination (CG) and infection stages (1.5, 3, 6, 12 and 24 h [IF1.5–IF24 h]) using reverse transcription‐quantitative PCR (RT‐qPCR). The results showed that PlRab8A was up‐regulated during the zoospore and cyst stages (Figure 1b), while PlRab8B was up‐regulated during the sporangial, zoospore, cyst and cyst germination stages and during IF1.5 h and IF3 h (Figure 1c).

2.2. Functional Characterisation of PlRab8A and PlRab8B

2.2.1. PlRab8A and PlRab8B Are Important for Mycelial Growth in P. litchii

To further determine the biological functions of PlRab8A and PlRab8B, we knocked out each of these two genes in the P. litchii wild‐type (WT) strain SHS3 using polyethylene glycol (PEG)‐mediated CRISPR/Cas9 gene‐editing technology (Figure 1d). Three PlRab8A knockout mutants (∆plrab8a: T39, T50 and T54) and three PlRab8B knockout mutants (∆plrab8b: T12, T16 and T33) were confirmed by PCR and sequencing. Strains that were subjected to transformation but in which knockout failed were used as control (CK) strains (Figure 1e).

To complement PlRab8A and PlRab8B mutants, PlRab8A‐GFP, RFP‐PlRab8B and PlRab8B‐GFP were inserted into the original genome position of PlRab8A and PlRab8B by CRISPR/Cas9. The complemented strains were named C96 (PlRab8A‐GFP), CR75 (RFP‐PlRab8B) and C36G (PlRab8B‐GFP) (Figure S2a,b). Microscopic observation revealed distinct subcellular localisation patterns between PlRab8A and PlRab8B. PlRab8A and PlRab8B exhibited both localised to membrane compartments, while PlRab8B additionally targeted distinct structures (subsequently identified as lipid droplets) that were absent in PlRab8A localisation patterns (Figure S2c). The RT‐qPCR analysis further revealed that the transcriptional levels of the corresponding genes in the complemented strains ∆plrab8a‐C96, ∆plrab8b‐C36G and ∆plrab8b‐CR75 reached or exceeded those of WT (Figure S2d). We measured the growth of these complemented strains 6 days post‐inoculation (dpi) on carrot juice agar (CJA) (Figure 2a). The results showed that the growth rates of the ∆plrab8a and ∆plrab8b were significantly lower compared to WT and complemented strains (Figure 2b), indicating that PlRab8A and PlRab8B contributed to the vegetative growth of P. litchii.

FIGURE 2.

The knockout of PlRab8A or PlRab8B affects the growth, sporangial morphology, sporangia production, zoospore release and oospores formation. (a) Colony morphology and growth of Peronophythora litchii wild‐type (WT), CKPlRab8 mutants (T39, PlRab8A knockout; T33, PlRab8B knockout) and complemented strains (C96, PlRab8A‐GFP; CR75, RFP‐PlRab8B; C36G, PlRab8B‐GFP), growing on carrot juice agar (CJA), photographs were taken 6 days post‐inoculation. (b) The chart of the growth rate of (a). (c) Statistical analyses of the number of sporangia in WT and PlRab8 mutants. CK, negative control. (d) Statistical analyses of sporangial length and width in WT and PlRab8 mutants. Sporangia from WT and PlRab8 mutants grown on CJA were placed into water and observed and measured for their length and width under a microscope. (e) Statistical analysis of zoospore release rates for WT and PlRab8 mutants. Data are mean ± SD. (f) The sporangia of WT and PlRab8 mutants were stained with DAPI (nuclei) or FM4‐64 (cell membranes), and then photographed under fluorescence. Sporangia were viewed under differential interference contrast (DIC). Bars = 10 μm. (g, h) Statistical analysis and microscopic observation of oospore formation in WT and PlRab8 mutants. Data are mean ± SD. (h) Microscopic observation of oospores in WT and PlRab8 mutants; blue arrows indicate the location of oospores. Bars = 100 μm. The experiments were independently repeated three times. Asterisks represent significant differences (t test, **p < 0.01; ***p < 0.001, ****p < 0.0001).

2.2.2. PlRab8A and PlRab8B Influence the Production and Morphology of Sporangia of P. litchii

Peronophythora litchii is transmitted as sporangia or zoospores (Situ et al. 2022). We measured the sporangial production of WT, CK, Δplrab8a and Δplrab8b after inoculating them on CJA for 7 days. The results showed that the sporangial production of the Δplrab8a and Δplrab8b was significantly higher than that of WT or CK. Meanwhile, ∆plrab8a‐C96, ∆plrab8b‐C36 and ∆plrab8b‐CR75 had restored sporangial production of the corresponding mutants to the WT level (Figure 2c). We also measured the length and width of sporangia. Compared with WT and CK, the sporangia of the Δplrab8b showed longer lengths and similar widths, while the sporangia of the Δplrab8a did not show observable differences in sporangial morphology (Figure 2d). These results indicated that PlRab8B is involved in the sporangial development of P. litchii.

2.2.3. PlRab8B Influences the Formation of Zoospores in P. litchii

Zoospores are released after sporangial cleavage (Zhang et al. 2024). In this study, we found that the sporangia of Δplrab8b only released a few zoospores (Figure 2e), but the complemented strain Δplrab8b‐CR75 and WT released numerous zoospores. Microscopic observation showed that the Δplrab8b exhibited aberrant membranes of zoospores in sporangia (Figure 2f). In contrast, the sporangial cleavage and zoospore release rates were not affected by the knockout of PlRab8A (Figure 2e,f). These results suggested that PlRab8B plays a critical role in the formation of the zoospore membrane in P. litchii.

2.2.4. PlRab8A and PlRab8B Are Involved in Cell Wall Integrity, and Osmotic Stress Tolerance, and Oospores Formation of P. litchii

Oospores exhibit greater environmental resistance, survive longer in the soil and often serve as the primary inoculum in the following year (Situ et al. 2022; Zhu et al. 2022). Here, we found that the oospore production of Δplrab8b was significantly decreased, while the oospore numbers of Δplrab8a significantly increased compared to WT and CK (Figure 2g,h). These results suggest that PlRab8B positively contributes to the production of oospores, while PlRab8A negatively contributes to the production of oospores in P. litchii.

We also tested the cell wall integrity of the above mutants using disruptors (Calcofluor white [CFW], Congo red [CR] and sodium dodecyl sulphate [SDS]) and osmotic stress agents (NaCl, CaCl₂ and sorbitol). These compounds were added separately to the Plich medium for testing. The growth inhibition of Δplrab8a was significantly lower than that of WT on medium containing CR, but there were no observable growth rate differences in response to SDS or CFW treatment (Figure 3a and Figure S3a). The growth inhibition of Δplrab8b was significantly lower than that of WT on media containing CR or CFW, but showed no growth differences in response to SDS treatment (Figure 3b and Figure S3b). These results indicated that these two genes are involved in resistance to cell wall stresses. In addition, Δplrab8a was more sensitive to sorbitol, NaCl and CaCl₂ compared to WT and CK (Figure 3a and Figure S3a). Δplrab8b also showed increased sensitivity to sorbitol and CaCl₂, but no significant differences were observed in Plich medium supplemented with NaCl compared to WT and CK (Figure 3b and Figure S3b). The complemented strains showed restored phenotypes under various stress conditions (Figure 3a,b). These results suggested that PlRab8A and PlRab8B may both be involved in osmotic stress tolerance.

FIGURE 3.

The knockout of PlRab8 affects cell wall integrity and osmotic stress tolerance. (a and b) Inhibition rate for Peronophythora litchii wild type (WT), PlRab8 mutants (PlRab8A: T39, T50, T54; PlRab8B: T12, T16, T33), and complemented strains (C96, PlRab8A‐GFP; C36G, PlRab8B‐GFP; CR75, RFP‐PlRab8B) as shown in Figure S3a,b, compared with WT and negative control (CK). WT and PlRab8 mutants were separately inoculated onto Plich medium supplemented with 20 μg·mL⁻¹ SDS, 350 μg·mL⁻¹ Congo red (CR), 100 μg·mL⁻¹ calcofluor white (CFW), 0.2 M sorbitol, 0.05 M NaCl or 0.1 M CaCl ₂, and then incubated at 25°C in the dark. Photographs were taken 6 days post‐inoculation. Data are mean ± SD based on three independent replicate experiments. Asterisks represent significant differences (t test, **p < 0.01; ***p < 0.001).

2.2.5. PlRab8A and PlRab8B Are Important for Full Virulence of P. litchii and Laccase Activity

To investigate the roles of PlRab8A and PlRab8B in the pathogenesis of P. litchii, hyphal plugs, zoospores and sporangial suspensions of WT and PlRab8 mutants were separately inoculated onto uninjured detached litchi leaves (Figure 4a,b). The lesion diameters caused by these mutants were significantly smaller than those caused by WT and the complemented strains at 48 h post‐inoculation (hpi) (Figure 4a–d). Laccase activity assay also showed that Δplrab8a and Δplrab8b exhibited decreased laccase activity (Figure 4e,f). These results demonstrated that PlRab8A and PlRab8B are associated with the pathogenicity and laccase activity of P. litchii.

FIGURE 4.

PlRab8A and PlRab8b are associated with the virulence and laccase activity of Peronophythora litchii. (a, b) The litchi leaves were inoculated with hyphal plugs or zoospores of wild‐type (WT) and mutants (T39, PlRab8A knockout; T33, PlRab8B knockout) and complemented strains (C96, PlRab8A‐GFP; CR75, RFP‐PlRab8B; C36G, PlRab8B‐GFP). Photographs were taken 48 h post‐inoculation. (c, d) Δplrab8a and Δplrab8b induced smaller lesions on lichi leaves. Data are mean ± SD. (e) WT and PlRab8 mutants were inoculated on lima bean supplemented with ABTS for testing laccase activity. (f) The statistical diagram shows the diameters of oxidised ABTS for the WT and PlRab8 mutants. Data are mean ± SD. The experiments were independently repeated three times. Asterisks represent significant difference (t test, **p < 0.01; ***p < 0.001).

Further analyses showed that the laccase activity and pathogenicity of CR75 (RFP‐PlRab8B) had recovered to WT level (Figure 4). However, the laccase activity and virulence of C36G (PlRab8B‐GFP) recovered to only 50% of WT (Figure 4) suggesting that the C‐terminal GFP tag may disrupt the functions of PlRab8B. These results suggested that the N‐terminal fluorescent protein tag was more suitable for PlRab8B.

2.3. The Role of PlRab8 in Lipid Droplet Regulation and Autophagy

2.3.1. Lipid Droplet Accumulation Is Regulated by PlRab8B

Because Rab proteins are integral components of organelle membranes, we used Nile Red stain on Δplrab8a, Δplrab8b and WT to examine lipid distribution. After cultivating Δplrab8a, Δplrab8b and WT on CJA medium for 5 days, we placed peripheral hyphae onto slides and observed that there was more unstained intracellular amorphous substance in Δplrab8b‐T33 than in WT. We added 2 μL of Nile Red stain to each slide. While Δplrab8a‐T50 showed no detectable change, the lipid droplets (LDs) within the hyphae of Δplrab8b‐T33 were significantly more abundant compared to those in WT hyphae (Figure 5a).

FIGURE 5.

The knockout of PlRab8B promotes the accumulation of lipids and enhances cellular autophagy. (a) Nile Red‐stained lipid vesicles and cellular contents observed under fluorescent microscope. Δplrab8b‐T33 and Peronophythora litchii wild type (WT) were cultured on carrot juice agar for 5 days and then stained with Nile Red. Bars = 25 μm. (b) The hyphal cells of WT and Δplrab8b‐T33 were observed by transmission electron microscope, the blue arrows indicate lipid droplets (LDs). Bars = 1 μm. (c) Boron‐dipyrromethene (BODIPY) staining indicated the localisation of LDs, which are partially co‐localised with RFP‐PlRab8B. Bars = 5 μm. (d) WT and Δplrab8b were cultured in Plich liquid medium for 5 days. Subsequently, the mycelia were rinsed three times with double‐distilled water (ddH₂O) and then treated separately in Plich liquid medium (CK) or ddH₂O (starvation treatment) for 4 h. These mycelia were stained with monodansylcadaverine (MDC) for 2 min in the dark and then rinsed with sterile distilled water three times, and photographed. Bars = 10 μm. (e) Transcriptional analysis of PlATGs in Δplrab8b‐T33 and WT. Data are mean ± SD based on three independent replicate experiments. Asterisks represent significant differences (t test, **p < 0.01; ***p < 0.001).

Transmission electron microscopic observation of hyphae showed that more LDs were accumulated in Δplrab8b‐T33 cells compared to WT (Figure 5b). Therefore, we wondered whether PlRab8B is associated with LDs. LD‐BODIPY (boron‐dipyrromethene) staining was used for visualising LDs in CR75. The results indicated that LD‐BODIPY colocalised with RFP‐PlRab8B (Figure 5c).

2.3.2. Knockout of PlRab8 Led to More Intense Autophagy in P. litchii

Rab proteins play key roles in intracellular cargo sorting, and intracellular autophagy requires accurate protein sorting (Bhuin and Roy 2014). Through monodansylcadaverine (MDC) staining, we found that there were more acidic vesicles in Δplrab8b‐T33, indicating that membrane transport was disturbed in this mutant. After mycelial starvation treatments, acidic vesicles increased in WT, while there was no significant change in Δplrab8b (Figure 5d).

These results demonstrated that the knockout of PlRab8B enhanced cellular autophagy. To further confirm the enhanced autophagy in Δplrab8a and Δplrab8b, we performed RT‐qPCR analysis on cDNA from Δplrab8b‐T33, Δplrab8a‐T50 and WT samples targeting 21 PlATGs. The results showed that the expression levels of most PlATGs were higher in Δplrab8a‐T50 and Δplrab8b‐T33 compared to the WT (Figure 5e and Figure S3c), suggesting that the knockout of PlRab8 led to enhanced autophagy in P. litchii.

2.4. PlRab8A Interacted With PlRab6

Rab8A‐GFP and Rab8B‐GFP proteins were extracted from P. litchii, and the interaction between PlRab6 and PlRab8A was found through co‐immunoprecipitation mass spectrometry (Co‐IP‐MS) experiments (Figure S4a). To further validate this interaction, we overexpressed only PlRab6‐RFP or RFP in Δplrab8a‐C96. Proteins were extracted and subjected to co‐immunoprecipitation (Co‐IP) assays using GFP‐trap beads. The results showed that PlRab8A‐GFP could effectively co‐isolate PlRab6‐RFP, confirming their interaction in vivo (Figure 6a). Additionally, we conducted in vitro glutathione S‐tranferase (GST) pull‐down assays. GST‐PlRab6 and His‐PlRab8A were expressed and purified. The GST pull‐down results indicated that GST alone could not pull down His‐PlRab8A, whereas GST‐PlRab6 successfully pulled down His‐PlRab8A, further supporting the direct interaction between PlRab6 and PlRab8A (Figure 6b). These results indicated that PlRab8A directly interacts with PlRab6.

FIGURE 6.

PlRab6 interacted with PlRab8A and is required for growth, zoospore release and pathogenicity of Peronophythora litchii. (a) Interaction between PlRab8A and PlRab6 in vivo. Total proteins were extracted from the hyphae of P. litchii expressing Rab8A‐GFP and Rab6‐RFP. Protein complexes were pulled down using GFP‐Trap‐A magnetic beads, and the co‐precipitation of these proteins was detected by western blot. (b) In vitro interaction between Rab8A and Rab6. His‐Rab8A and GST‐Rab6 were expressed in Escherichia coli . The co‐precipitation of His‐Rab8A and GST‐Rab6 was detected by western blotting after affinity purification using GST‐agarose beads. (c) Transcription levels of PlRab6 during mycelial (MY), sporangium (SP), zoospore (ZO), cystospore (CY), cyst germination (CG) and 1.5 to 24 h post‐infection (IF1.5–24 h). The P. litchii actin gene was used as internal control, and results were calculated using the 2^−ΔΔCt method. Data are mean ± SD based on three independent replicate experiments. (d) Growth rates for wild type (WT), control (CK), Δplrab6 (T2, T21 and T77). Data are mean ± SD. (e) Growth inhibition for Δplrab6 compared with WT and CK, under stress conditions of 50 μg·mL⁻¹ SDS, 350 μg·mL⁻¹ Congo red (CR), 100 μg·mL⁻¹ calcofluor white (CFW), 0.2 M sorbitol, 0.05 M NaCl or 0.1 M CaCl₂. Data are mean ± SD based on three independent replicate experiments. (f) Sporangial production of WT, CK, T2, T21 and T77. Data are mean ± SD. (g) Oospore production of WT, CK, T2, T21 and T77. Data are mean ± SD. (h) The release rate of zoospores in WT, CK, T2, T21 and T77. Data are mean ± SD. (i) The sporangia of WT and Δplrab6 were separately stained with DAPI and FM4‐64, followed by microscopic observation under a fluorescence microscope to visualise the nuclei and membranes. These experiments were independently repeated three times. Black asterisks represent significant differences (t test, **p < 0.01; ***p < 0.001). Red asterisks indicate the protein bands of correct size.

2.5. PlRab6 Is Required for Growth, Zoospores Formation and Pathogenicity of P. litchii

Because of the observed interactions between PlRab8A and PlRab6, we further investigated the function of PlRab6. The CDS region of the PlRab6 gene was 546 bp in length, encoding 182 amino acids. Transcriptional analysis revealed that PlRab6 expression was up‐regulated during the zoospore stage, cyst stage and IF1.5 h (Figure 6c), suggesting a potential role at these stages. To further explore the function of PlRab6, we used CRISPR/Cas9 gene‐editing technology to knock out PlRab6 and confirmed the knockout of PlRab6 in mutants (∆plrab6: T2, T21, T77) through PCR and sequencing (Figure S4b). We inoculated hyphal plugs of ∆plrab6, WT and CK onto CJA and measured the colony diameter 6 dpi (Figure S4c). The results showed that the mycelial growth rate of Δplrab6 was significantly reduced compared to WT and CK (Figure 6d). We also tested the cell wall integrity and osmotic stress tolerance of Δplrab6. Δplrab6 showed less sensitivity against 350 μg·mL⁻¹ CR and 50 μg·mL⁻¹ SDS compared to WT (Figure 6e). Under conditions of 0.2 M sorbitol and 0.1 M CaCl₂, the growth inhibition of Δplrab6 was significantly lower than that of the WT, whereas at 0.05 M NaCl, the colony diameter of Δplrab6 was slightly larger than that on the non‐amended Plich medium, with indications of growth stimulation (negative growth inhibition) being displayed (Figure 6e and Figure S4d). Knockout of PlRab6 did not affect the tolerance of P. litchii against CFW. These results indicated that PlRab6 is associated with the tolerance of P. litchii to cell wall and osmotic stress.

Knockout of PlRab6 did not affect the production of sporangia and oospores (Figure 6f,g). However, zoospore release rates of Δplrab6 were significantly decreased (Figure 6h). DAPI and FM4‐64 staining revealed that the formation of normal cell nuclei within sporangia was hindered in Δplrab6 (Figure 6i). These results indicated that PlRab6 regulates the zoospore formation via involvement of the nucleus formation in sporangia.

For pathogenesis assays, hyphal plugs or zoospore suspensions each of T2, T21, T77, WT and CK were inoculated onto detached litchi leaves, and lesion lengths were measured at 48 hpi (Figure S5a). Statistical analysis showed that lesions caused by Δplrab6 were significantly smaller than those of WT and CK (Figure 7a,b). These findings demonstrated that PlRab6 is crucial for the pathogenicity of P. litchii. Additionally, we tested the laccase activity of Δplrab6 and WT. Δplrab6 produced less oxidised ABTS [2,2′‐Azino‐bis (3‐ethylbenzothiazoline‐6‐sulfonic acid) diammonium salt] compared to WT (Figure S5b). The statistical analysis revealed significantly decreased laccase activity in Δplrab6 compared to WT (Figure 7c). To further investigate the role of PlRab6 in cellular processes, we analysed the transcriptional levels of PlATGs in Δplrab6‐T21 and WT samples. The results showed that the expression levels of most PlATGs were significantly up‐regulated in Δplrab6‐T21 compared to WT (Figure S5c). This suggested that the knockout of PlRab6 may lead to an enhancement of autophagy in P. litchii, potentially as a compensatory mechanism for the loss of PlRab6 function.

FIGURE 7.

PlRab6 is required for the full virulence and laccase activity of Peronophythora litchii. (a) Pathogenicity assays of Δplrab6 on litchi leaves. The hyphal plugs and one hundred zoospores from wild type (WT), control (CK), gene knockout mutants (T2, T21 and T77) were inoculated on the uninjured detached litchi leaves. Photographs were taken at 48 h post‐inoculation (hpi). (b) Lesion diameters of P. litchii infection at 48 hpi. Data are mean ± SD. (c) Laccase activity assay. WT and Δplrab6 were inoculated on lima bean medium supplemented with ABTS for 6 days, the diameter of oxidised ABTS were measured 6 days post‐inoculation. The experiments were repeated three times. Asterisks represent significant differences (t test, ***p < 0.001). (d) A schematic diagram illustrating the interaction and functions of Rab proteins (Rab6, Rab8A and Rab8B) from P. litchii. The three Rab proteins were involved in regulating the growth, pathogenicity, lipid droplet distribution and autophagy of P. litchii.

Taken together, PlRab6 interacts with PlRab8A, and both proteins regulate the growth, cell wall integrity, osmotic tolerance, zoospore formation, pathogenicity and laccase activity in P. litchii. However, we also observed functional differences between these two proteins. Specifically, in Δplrab8a‐T50, the transcriptional levels of both PlRab6 and PlRab8B were significantly up‐regulated (Figure S5d). Conversely, in Δplrab6‐T21, the transcriptional level of PlRab8A was markedly down‐regulated (Figure S5d). These findings suggested that PlRab6 and PlRab8A may have distinct roles in the regulatory network of P. litchii, and their interactions and expression patterns are intricately linked to the cellular functions and pathogenicity of the organism.

3. Discussion

Rab GTPases are known to regulate diverse cellular processes, including hyphal growth, apical polar growth, colony morphology, conidiation, autophagy, pathogenicity, protein trafficking and secretion (Bielska et al. 2014; Irieda et al. 2014; Liu et al. 2015). However, the functions of Rab proteins in oomycetes remain poorly understood, limiting our ability to target these pathways for disease control.

In this study, we identified 36 putative Rab genes in P. litchii that showed various transcription profiles (Figure S1). Notably, PlRab8A was up‐regulated during zoospore and cyst stages, while PlRab8B showed increased expression in sporangia, zoospores, cysts, during cyst germination and at early infection stages (Figure 1b,c), suggesting their roles in asexual reproduction and pathogenicity. Consistent with findings in F. graminearum (FgRab8) and M. oryzae (MoSec4), knockout of PlRab8 significantly impaired the growth and pathogenicity of P. litchii (Figures 2a,b and 4) (Zheng et al. 2015, 2016). SEC4/Rab8, as the first Rab protein discovered in yeast, has been found to be associated with protein secretion in yeast cells (Liu et al. 2015). Furthermore, the knockout of PlRab8 resulted in reduced laccase activity (Figure 4e,f), indicating that PlRab8 may regulate virulence through protein secretion and growth modulation.

Interestingly, ∆plrab8a and ∆plrab8b exhibited increased sporangial production compared to WT (Figure 2d), with ∆plrab8b sporangia being significantly longer (Figure 2e). However, zoospore release was dramatically impaired in ∆plrab8b (Figure 2e), likely due to defects in vesicle trafficking and membrane remodelling, which are critical for sporangial cleavage and flagella assembly (Judelson and Blanco 2005). Notably, the ∆plrab6 exhibited reduced zoospore release compared to WT, likely caused by nuclear integrity defects specific to these mutants (Figure 6h,i). This suggests that the underlying mechanism for impaired zoospore release differs from that in the ∆plrab8b. These findings align with previous reports on PlMAPK2 and PsMAD1, which are also essential for sporangial cleavage (Lin et al. 2018; Huang et al. 2021). While PlMAPK10 has been shown to positively regulate sporangial production (Jiang et al. 2018), the precise relationships on sporangial cleavage and production between these proteins remain unclear, highlighting the need for further research to elucidate the asexual development network in P. litchii. Oospores, the overwintering structures of P. litchii, were produced in significantly higher quantities in ∆plrab8a but reduced in ∆plrab8b, suggesting divergent roles of these Rab proteins in sexual reproduction. Previous studies have indicated that PlM90 and the C2H2 zinc finger protein CZF1 are associated with oospore development (Jiang et al. 2017; Zhu et al. 2022). But the functional relationships between these proteins and Rab GTPases remain to be explored.

In mice, Rab8a and Rab6 exhibit co‐localisation and functional interplay (Grigoriev et al. 2011). Here, we provide the first evidence of an interaction between PlRab8A and PlRab6 in oomycetes (Figure 6a,b). The ∆plrab6 and ∆plrab8 both showed reduced growth, pathogenicity and stress tolerance. Rab proteins have repeatedly been implicated in cilia‐related functions (Follit et al. 2010; Zhang et al. 2015; Aljiboury et al. 2023). Additionally, both ∆plrab6 and ∆plrab8 exhibited impaired zoospore release. Collectively, these findings indicated that PlRab8 and PlRab6 may, in coordination, affect growth, cell wall integrity, osmotic tolerance and pathogenicity in P. litchii.

Rab8a is known to mediate LD–mitochondria interactions, facilitating the transfer of long‐chain fatty acids (Ouyang et al. 2023). Our results demonstrate that PlRab8B is involved in autophagy and LD accumulation in P. litchii. Knockout of PlRab8 and PlRab6 led to the up‐regulation of most PlATGs (Figure 5e; Figures S3c and S5c). The knockout of PlRab8B altered LD dynamics (Figure 5a,b). These findings, combined with previous reports on PlATG6a and PlATG8 (Wang et al. 2022; Yu et al. 2024), suggest that PlRab8B may regulate development and pathogenesis through lipid metabolism and modulation of autophagy, which is a novel insight into Rab GTPase function in oomycetes.

Despite their importance, the functions of Rab GTPases in oomycetes remain poorly characterised. This study demonstrates that PlRab8A, PlRab8B and PlRab6 are critical for growth, sexual and asexual development, and pathogenicity in P. litchii. Specifically, PlRab6 and PlRab8A may form a functional complex regulating growth, virulence and autophagy, while PlRab8B likely modulates sporangial development and pathogenicity through autophagy and LD metabolism (Figure 7d). Future studies should explore how Rab proteins coordinate intracellular membrane trafficking, autophagy and lipid metabolism to regulate development and pathogenicity.

4. Experimental Procedures

4.1. Strains and Sequences

The P. litchii wild‐type strain SHS3 (WT) was isolated and identified from diseased leaves of litchi trees in this laboratory. The CK (control strain) was an attempted transformant that failed to knock out PlRab8 or PlRab6. P. litchii WT, CK, Δplrab8a, Δplrab8b and Δplrab6 were maintained on CJA (juice from 250 g carrot tissue for 1 L medium, with agar 15 g·L⁻¹ for solid media) at 25°C in the dark. Escherichia coli DH5α was cultured in Luria Bertani (LB) medium at 37°C for the cloning and propagation of recombinant plasmids.

The genomic sequence of P. litchii was obtained from NCBI (BioProject ID: PRJNA290406) (Ye et al. 2016).

4.2. Bioinformatic Analysis

TBtools was used to retrieve the protein sequence of PlRab8 from a database specific to P. litchii (Chen et al. 2023). After BLAST against the NCBI database (https://www.ncbi.nlm.nih.gov/), homologues of Rab8 were obtained (including Ph. fragariae, Ph. ramorum, Ph. boehmeriae, Py. oligandrum and Py. brassicum). Sequences were submitted to NCBI‐CDD (http://www.ncbi.nlm.nih.gov/Structure/cdd) and SMART (https://smart.embl.de/) to identify conserved domains. Phylogenetic analysis was conducted using MEGA 11 software with the neighbour‐joining algorithm for cluster analysis. For analysis of consistent branching, bootstrap replications were repeated 1000 times.

4.3. RT‐qPCR Analysis

Total RNA was isolated with the RNA Extraction Kit (Sanger) and reverse transcribed into complementary DNAs (cDNAs) with a FastQuant RT Kit (Takara). Each cDNA sample was used as a template to determine transcriptional levels of P. litchii GTPase‐encoding genes via qPCR with specific primer pairs (Table S1) using actin as an internal control. The relative transcriptional level of each gene was computed using the 2^−ΔΔCt method. The primers used in this study are listed in Table S1.

4.4. Plasmid Construction

The PCR fragments were amplified using Phanta Max Super‐Fidelity DNA Polymerase (Vazyme). PlRab genes were amplified from the cDNA of WT. The amplified fragments of PlRab6 and PlRab8A were digested with BamHI and EcoRI, respectively, and cloned into pET32a and pGEX‐6P‐1 for protein expression and purification in E. coli . The vectors pYF2.3G‐RibosgRNA and pBluescript II KS used for the knockout of PlRab by CRISPR/Cas9 were generated as described previously (Fang and Tyler 2016).

4.5. CRISPR/Cas9 Editing for Gene Knockout

Peronophythora litchii was transformed to delete PlRab genes as previously described (Fang and Tyler 2016). The targets of sgRNA1 and sgRNA2 are shown in Figure 1d. Potential off‐target sites were examined using the FungiDB alignment search tool (BLASTN) against the P. litchii genome. sgRNA1 and sgRNA2 were inserted into the sgRNA vector pYF2.3G‐RibosgRNA, which had been cleaved by NheI and BsaI. Two sgRNA vectors (PlRab genes), pBSSK (PlRab genes) and the hSPCas9 vector pYF2‐PsNLS‐hSpCas9, were co‐transformed into P. litchii using PEG‐mediated protoplast transformation. The transformants were screened using G418, PCR and sequencing assays.

4.6. Analyses of Sporangial, Zoospore and Oospore Development

For sporangial production, five hyphal plugs were obtained with a 9‐mm‐diameter cork borer from the growing edge of a 5‐day‐old colony and placed in 5 mL of water to obtain sporangia. Zoospores were obtained by incubating under cold shock at 4°C for 20 min. For oospore production, P. litchii were inoculated on CJA and incubated at 25°C in the dark for 15 days. Subsequently, a single hyphal plug (2 × 2 cm) from near the inoculation point was homogenised in 5 mL of water, and an aliquot of 2 μL of the mixture was assessed microscopically for oospore production.

To visualise the distribution of nuclei, sporangia were stained with 4′,6‐diamidino‐2‐phenylindole (DAPI; Beyotime). To investigate the sporangial cleavage, we stained the sporangia using FM 4‐64 dye (N‐(3‐triethylammoniumpropyl)‐4‐(6‐(4‐(diethyla‐mino) phenyl) hexatrienyl) pyridinium dibromide; Invitrogen), which binds to the plasma membrane.

4.7. Autophagy Analysis

To investigate whether the knockout of PlRab8B affected autophagy in P. litchii, we cultured both WT and Δplrab8b in Plich liquid medium for 5 days. Subsequently, the mycelia were rinsed three times with double‐distilled water and then treated separately in Plich medium supplemented with 2 mM phenylmethylsulfonyl fluoride (PMSF) and double‐distilled water (starvation treatment) for 4 h. Fresh mycelia were then stained with 2 μL monodansylcadaverine (MDC; Yuanye) in the dark for 2 min, and then rinsed with sterile distilled water three times. The samples were visualised by a fluorescent microscope.

4.8. Stress Tolerance Tests

The hyphal plugs of each strain were inoculated onto Plich agar supplemented with 350 μg·mL⁻¹ Congo red (CR), 100 μg·mL⁻¹ calcofluor white (CFW), 20 μg·mL⁻¹ SDS, 0.1 M CaCl₂, 0.05 M NaCl or 0.2 M sorbitol and incubated at 25°C for 6 days. Then, the growth rate was measured. The growth inhibition was calculated from diameters on amended and non‐amended Plich media where inhibition = (non‐amended growth − amended growth)/non‐amended growth.

4.9. Pathogenicity Assays

For pathogenicity assays, zoospores or sporangial suspensions were inoculated onto detached tender leaves of litchi (cv. Guiwei), which were collected from the litchi orchard at the South China Agricultural University, Guangzhou, Guangdong province. One hundred zoospores or sporangia of each strain were inoculated onto the centre of each leaf and kept at 80% humidity at 25°C in the dark. The lesion diameters were measured at 48 hpi. The experiments were repeated three times, with each repetition consisting of 10 technical replicates.

4.10. Protein Extraction and Western Blot Analysis

PEG‐mediated transformation and CRISPR/Cas9 technology were used to reintroduce PlRab8A‐GFP to its natural locus in knockout mutants. Following transformation, the resulting mutants were selected and grown in carrot juice broth for 7 days at 25°C. Then the total proteins were extracted from mycelia with RIPA buffer (Thermo Fisher Scientific). The presence of the proteins was detected by anti‐GFP antibodies (Abmart) as previously described (Li et al. 2024).

4.11. Co‐Immunoprecipitation Assay

The plasmids pTOR‐RFP and pTOR‐Rab6‐RFP were introduced into the transformant expressing C96. Total proteins of the transformants expressing Rab8A‐GFP and RFP‐PlRab6 were extracted with RIPA buffer. The extracted proteins were incubated with 20 μL GFP magnetic beads for 1 h at 4°C. Then these beads were washed three times with lysis buffer. The presence of Rab8A‐GFP and RFP‐PlRab6 was detected by western blot; RFP was used as control.

4.12. GST Pull‐Down Assay

The pGEX‐6p‐1, pGEX‐6p‐1‐PlRab6 and pET32a‐His‐PlRab8A plasmids were transformed into E. coli BL21 to produce GST, GST‐tagged PlRab6 and His‐tagged PlRab8A proteins. E. coli BL21 was incubated for 12 h at 22°C in LB amended with 0.1 mM isopropyl‐β‐D‐thiogalactopyranoside (IPTG). These bacteria were pelleted and lysed using a sonic dismembrator in cold lysis buffer (phosphate‐buffered saline, 1% vol/vol Triton X‐100% and 1% vol/vol phenylmethylsulfonyl fluoride). GST and GST‐PlRab6 were incubated with 40 μL GST magnetic beads (Thermo Fisher Scientific) for 1 h at 4°C. Then these beads were washed three times with lysis buffer. Next, the beads were incubated with His‐PlRab8A for 1 h at 4°C, washed three times and then boiled for 10 min. The presence of GST‐ or His‐tagged proteins was assessed using western blot analysis.

Author Contributions

Gangqiang Zhou: data curation, methodology, investigation, software, validation, visualization, writing – original draft. Honglin Chen: software and data curation. Wenzhe Zhou, Rubin Shen, Gao Gao and Lizhu Xie: data curation. Junjian Situ, Pinggen Xi, Minhui Li, Wen Li, Tom Hsiang and Zide jiang: writing – review and editing. Guanghui Kong: conceptualization, data curation, writing – review and editing, supervision. All authors have read and agreed to the published version of the manuscript.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Identification and Transcriptional profile of Rab proteins in P. litchii. In the cluster tree, branches highlighted in red represent proteins predicted by NCBI CDD to contain RAB domains. The designations Plitchii008012.m1 and Plitchii004269.m1 correspond to PlRab8A and PlRab8B. In the graph, the redder the colour, the higher the transcription level of these genes during the life cycle of P. litchii. Neighbour‐joining analysis using MEGA 11 with 1000 bootstrap replicates shown as percentages along branches.

Figure S2: Knockout and complementation of PlRab8A and PlRab8B, along with their subcellular localization analysis in P. litchii. (a) PCR and electrophoresis analyses confirmed the knockout and complementation of PlRab8A and PlRab8B in P. litchii. (b) subcellular localization analysis PlRab8A and PlRab8B in P. litchii. (c) Transcriptional analysis of PlRab8A and PlRab8B gene in WT and complemented strains. Data are mean ± SD based on three independent replicate experiments. Asterisks represent significant differences (t test, **p < 0.01; ***p < 0.001).

Figure S3: The knockout of PlRab8 affects cell wall integrity, osmotic stress tolerance and enhances cellular autophagy. (a and b) The sensitivity of WT and PlRab8 mutants to cell wall integrity and osmotic stress was examined. WT and PlRab8 mutants were separately inoculated onto Plich medium supplemented with 20 μg·mL⁻¹ SDS, 350 μg·mL⁻¹ CR, 100 μg·mL⁻¹ CFW, 0.2 mol·L⁻¹ sorbitol, 0.05 mol·L⁻¹ NaCl or 0.1 mol·L⁻¹ CaCl₂, and then incubated at 25°C in the dark. Photographs were taken 6 dpi. (c) Transcriptional analysis of PlATGs in Δplrab8a‐T50 and WT. Data are mean ± SD based on three independent replicate experiments. Asterisks represent significant differences (t test, **p < 0.01; ***p < 0.001).

Figure S4: The knockout of PlRab6 affects growth. (a) PlRab8A‐GFP Co‐IP‐MS mass spectrometry analyses of PlRab8A‐interacting proteins. (b) CRISPR/Cas9‐mediated knockout of the PlRab6 gene and PCR analysis of Δplrab6 (T2, T21, T77) in P. litchii. (c) The colony morphology of WT, CK, Δplrab6. WT, CK and Δplrab6 were inoculated on CJA and incubated in the dark at 25°C, and photographs were taken 6 dpi. (d) WT, CK, Δplrab6 were separately inoculated onto non‐amended Plich medium or Plich medium supplemented with 50 μg·mL⁻¹ SDS, 350 μg·mL⁻¹ CR, 100 μg·mL⁻¹ CFW, 0.2 mol·L⁻¹ sorbitol, 0.05 mol·L⁻¹ NaCl or 0.1 mol·L⁻¹ CaCl₂. The cultures were then incubated in the dark at 25°C for 6 days before photography and growth assessments.

Figure S5: Knockout of PlRab6 affects zoospore release, and laccase activity of P. litchii. (a) Microscopic observation of zoospore release from sporangia in WT, CK and Δplrab6, including released sporangia, unreleased sporangia and zoospores. Bars = 100 μm. (b) Colony morphology and growth of WT and Δplrab6 grown in 15 mL of Lima bean medium supplemented with ABTS for 6 days. (c) Transcriptional analysis of PlATGs in Δplrab6‐T21 and WT. (d) Transcriptional analysis of PlRab6, PlRab8A and PlRab8B gene in WT, Δplrab6‐T21, Δplrab8a‐T50 and Δplrab8a‐T33. Data are mean ± SD based on three independent replicate experiments. Asterisks represent significant differences (t test, **p < 0.01; ***p < 0.001; ****p < 0.0001).

Table S1: Primers used in this study.

Zhou, G. , Chen H., Zhou W., et al. 2025. “Regulation of Oomycete Autophagy, Lipid Droplet Accumulation and Pathogenesis by Three Rab GTPases .” Molecular Plant Pathology 26, no. 9: e70154. 10.1111/mpp.70154.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.