Retromer protein VPS29 plays a crucial and positive role in the sumoylation system mediated by E3 SUMO ligase SIZ1

SUMMARY

Vacuolar protein sorting 29 (VPS29) functions in retrograde protein transport as a component of the retromer complex. However, the role of VPS29 in the regulation of post‐translational modifications, such as sumoylation and ubiquitination, has not been elucidated. In this study, we demonstrate that VPS29 positively regulates SIZ/PIAS‐type E3 SUMO (Small ubiquitin‐related modifier) ligase‐mediated sumoylation systems. In Arabidopsis, vps29‐3 mutants display upregulated salicylic acid (SA) signaling pathways and reactive oxygen species accumulation, similar to those observed in siz1 mutants. Arabidopsis VPS29 (AtVPS29) directly interacts with the Arabidopsis E3 SUMO ligase SIZ1 (AtSIZ1) and localizes not only to the cytoplasm but also to the nucleus. The loss of AtVPS29 leads to a depletion of AtSIZ1, whereas the E3 ubiquitin ligase constitutive photomorphogenic 1 (COP1), an upstream regulator of AtSIZ1, accumulates in vps29‐3 mutants. Conversely, overexpression of AtVPS29 results in the accumulation of AtSIZ1 and the depletion of COP1 in transgenic Arabidopsis. Similarly, in human cells, silencing of hVPS29 leads to the depletion of the E3 SUMO ligase, PIAS1, and the accumulation of huCOP1. Under heat stress conditions, the levels of SUMO‐conjugates are significantly lower in Arabidopsis vps29‐3 mutants, indicating a regulatory role of AtVPS29 on AtSIZ1 activity. Moreover, AtVPS29 inhibits ubiquitination pathway‐dependent degradation of AtSIZ1. Notably, AtSIZ1 forms a complex with AtVPS29 and trimeric retromer proteins. Taken together, our results indicate that VPS29 plays an essential role in signal transduction by regulating SIZ/PIAS‐type E3 ligase‐dependent sumoylation in both plants and animals.

Significance Statement

Vacuolar protein sorting 29 (VPS29) is known to function in retrograde protein transport as a component of the retromer complex. However, the role of VPS29 in the regulation of post‐translational modifications, including sumoylation, has not been elucidated. In this work, we provide, for the first time, evidence that VPS29 positively controls signal transduction through the sumoylation of target proteins via the activity of SIZ/PIAS‐type E3 SUMO ligases in both plant and animal systems.

INTRODUCTION

SUMO is a small polypeptide that is covalently attached to target proteins to modify their functions (Bayer et al., 1998). Sumoylation affects protein subcellular localization, protein function and stability, nuclear‐cytosolic transport, transcriptional regulation, apoptosis, stress responses, cell‐cycle progression, mitochondrial dynamics, and responses to DNA damage (Jmii & Cappadocia, 2021; Wilson, 2017).

SAP/Miz1 (SIZ1) is an E3 SUMO ligase that contains a Siz‐Protein Inhibitor of Activated STAT (PIAS) RING (SP‐RING), a chromatin organization domain known as SAF‐A/B‐Acinus‐PIAS (SAP), a valine‐proline (VP) motif, a SUMO‐interacting motif (SIM), and a nuclear localization signal (NLS) (Jackson, 2001; Jmii & Cappadocia, 2021). Plant SIZ1 contains an additional domain, the plant homeodomain (PHD). So far, three types of E3 SUMO ligases have been identified in plants: SIZ, PIAL, and HYPLOIDY2/MMS21. The SIZ type contains all domains and motifs; the PIAL type does not contain the SAP, PHD, VP1, VP2, and NLS domains; and MMS21 only contains SP‐RING (Jmii & Cappadocia, 2021).

SIZ1 and the SUMO machinery are present in the same nuclear bodies (NBs). The components of the SUMO machinery localize to NBs in a conjugation‐dependent manner (Mazur et al., 2018), while SIZ1 localizes to NBs in an SP‐RING‐dependent manner (Cheong et al., 2009). Numerous sumoylation substrates also localize to these NBs (Mazur et al., 2018). Constitutive photomorphogenic 1 (COP1) also localizes to the same NBs (Mazur et al., 2018), and its stability is regulated by the E3 SUMO ligase activity of AtSIZ1 (Lin et al., 2016). In turn, AtSIZ1 stability is regulated through its ubiquitination by the E3 ligase activity of COP1 (Kim et al., 2017; Kim, Jang, & Seo, 2016; Lin et al., 2016). Additionally, the nuclear localization of COP1, which is induced by various conditions, including light and temperature, leads to degradation of target proteins through its E3 ligase activity (Jia et al., 2014; Park et al., 2017; Tsuchiya et al., 2010; Yu et al., 2013). This indicates that ubiquitination of target proteins by COP1 occurs in the same NBs, given that COP1 localizes in speckles (Kim, Jang, & Seo, 2016; Mazur et al., 2018; Seo et al., 2003, 2004).

The E3 SUMO ligases in Arabidopsis thaliana, including AtSIZ1, HYPLOIDY 2/MMS21 and PIAL1 and PIAL2, are involved in several developmental processes, including seed germination (Kim, Kwak, et al., 2016), nutrient assimilation (Miura et al., 2005; Park et al., 2011), sulfur metabolism (Tomanov et al., 2014), hormone signaling (Kim et al., 2015; Zhang et al., 2019), and flowering (Jin et al., 2008; Miura et al., 2010; Son et al., 2014). In addition, plant E3 SUMO ligases play important roles in responses to abiotic stresses, such as low temperature (Miura et al., 2007), drought (Catala et al., 2007), heat (Kim, Jang, & Seo, 2016; Li et al., 2013; Zhang et al., 2018; Zhou et al., 2018), and high salinity (Miura et al., 2011; Tomanov et al., 2014). Recent studies have also indicated regulatory roles of AtSIZ1 in thermomorphogenesis (Hammoudi et al., 2018; Yu et al., 2024), plant immunity (Hammoudi et al., 2018; Niu et al., 2019), and photomorphogenesis (Xiong et al., 2023; Zhang et al., 2020).

Retromers are protein complexes composed of a large subunit consisting of vacuolar protein sorting (VPS) 26, VPS29, and VPS35, and a small subunit consisting of sorting nexin (SNX) dimers, VPS5 in yeast, and SNX1/2 in animals and plants. They were first characterized in yeast (Horazdovsky et al., 1997; Nothwehr & Hindes, 1997; Paravicini et al., 1992; Seaman et al., 1997). The retromer complex is a master regulator of endosomal dynamics, which is responsible for cargo sorting and recycling within tubulovesicular transport carriers (Seaman, 2021). Retromer mutation or dysregulation causes defective endosomal and lysosomal functions in humans, leading to neurodegenerative disorders, such as Parkinson's disease, Alzheimer's disease, and amyotrophic lateral sclerosis (Seaman, 2021).

VPS26, VPS29, and VPS35 proteins interact with each other in plants and colocalize in the prevacuolar compartment (PVC) (Oliviusson et al., 2006). These three retromer proteins are involved in various plant developmental processes, including programmed cell death (Jaillais et al., 2007; Munch et al., 2015; Zelazny et al., 2013), oil body biogenesis and breakdown during vegetative growth (Thazar‐Poulot et al., 2015), and root hair expansion (Jha et al., 2018). In particular, VPS29 participates in the maturation of storage proteins during seed development (Shimada et al., 2006) and the establishment of cell polarity during plant organogenesis (Jaillais et al., 2007). However, the exact molecular mechanisms by which retromer proteins regulate post‐translational modifications and are involved in developmental processes remain to be elucidated.

Arabidopsis thaliana siz1 mutants display severe dwarfism and low fertility, and their phenotypes are restored to wild‐type (WT) levels by exogenous supply of ammonium (Catala et al., 2007; Park et al., 2011). A. thaliana vps29‐3 mutants are also severely dwarfed and exhibit abnormal seed development (Jaillais et al., 2007; Shimada et al., 2006). Moreover, it was recently reported that AtVPS29 positively regulates the gibberellic acid (GA) signaling pathway, as vps29‐3 mutants harbor lower levels of SLEEPY1 (SLY1) and higher levels of REPRESSOR OF ga1‐3 (RGA) protein (Min et al., 2024). siz1‐2 mutants also exhibit similar protein levels because AtSIZ1 sumoylates and stabilizes SLY1, thereby activating GA signaling (Kim et al., 2015). These similarities between siz1 and vps29‐3 suggest that AtSIZ1‐mediated sumoylation may be impaired in vps29‐3 mutants.

In this study, we focused on the role of VPS29 protein in the regulation of SIZ/PIAS‐type E3 SUMO ligase‐dependent sumoylation of target proteins. We observed that COP1, an upstream regulator of AtSIZ1, accumulated in the vps29‐3 mutants, leading to a decrease in AtSIZ1 levels and a significant reduction in the sumoylation of target proteins. Similar results were obtained in human cells. The levels of E3 SUMO ligase PIAS1 decreased, and E3 ubiquitin ligase huCOP1 accumulated in hVPS29‐silenced HeLa cells. Additionally, we found that SUMO‐conjugates accumulated to a lesser extent in Arabidopsis vps29‐3 mutants, and that AtSIZ1 formed a complex with AtVPS29 and trimeric retromer proteins. Our results provide insights into how both plants and animals utilize VPS29 activity to modulate signaling mediated by the SIZ/PIAS‐type E3 SUMO ligase‐dependent sumoylation system.

RESULTS

SA signaling is upregulated in vps29‐3 mutants

In siz1 mutants, defects in the sumoylation system result in a severe dwarf phenotype (Catala et al., 2007), and vps29‐3 mutants also display severe dwarfism (Jaillais et al., 2007). Therefore, we investigated the phenotypic correlation between siz1 and vps29‐3 mutants. Previous studies have reported that Arabidopsis siz1 mutants accumulate higher levels of salicylic acid (SA) than WT plants, thereby exhibiting elevated reactive oxygen species (ROS) levels and increased immune responses (Castro et al., 2022; Lee et al., 2007). Thus, we extended our investigation to include the identification of SA signaling in vps29‐3 mutants. We first examined the transcript levels of SA signaling genes in vps29‐3 mutants. The results showed that the expression levels of the positive regulators of SA signaling, including ISOCHORISMATE SYNTHASE 1 (ICS1), ENHANCED DISEASE SUSCEPTIBILITY 5 (EDS5), PHYTOALEXIN DEFICIENT 4 (PAD4), and EDS1, were higher in vps29‐3 mutants than in the WT (Figure 1a), and these were previously shown to be upregulated in siz1 mutants (Lee et al., 2007).

Figure 1.

Salicylic acid (SA) signaling profiles of vps29‐3 mutants.

(a) Total RNA was isolated from 15‐day‐old wild‐type (WT), vps29‐3, and siz1‐2 plants. ICS1, EDS5, PAD4, and EDS1 transcript levels were examined using real‐time qRT‐PCR with each respective gene‐specific primer. Results are expressed as means ± SD (n = 3). Empty circles indicate individual data points. Asterisks indicate statistically significant differences in transcript levels (*P < 0.05; **P < 0.01; ***P < 0.001; Student's t‐test) between WT and vps29‐3 plants, or WT and siz1‐2 plants.

(b) Fifteen‐day‐old WT, vps29‐3, and siz1‐2 plants were subjected to histochemical H₂O₂ staining with 3,3′‐diaminobenzidine (DAB). Scale bar = 0.5 cm.

(c) Total RNA was isolated from the same samples described in (a). PR1 and PR2 transcript levels were examined using real‐time qRT‐PCR with PR1‐, and PR2‐specific primers. Results are expressed as means ± SD (n = 3). Empty circles indicate individual data points. Asterisks indicate statistically significant differences in transcript levels of PR1 and PR2 (**P < 0.01; ***P < 0.001; Student's t‐test) between WT and vps29‐3 plants, or WT and siz1‐2 plants.

Next, we analyzed ROS levels in vps29‐3 mutants, as ROS homeostasis is tightly connected to SA signaling (Herrera‐Vásquez et al., 2015). Our histochemical staining of H₂O₂ with 3,3′‐diaminobenzidine (DAB) revealed that vps29‐3 mutants accumulated substantially higher amounts of H₂O₂ than WT and even siz1‐2 mutants across the whole plants (Figure 1b). Specifically, while siz1‐2 mutants displayed more DAB staining than WT plants in the overall tissues of cotyledons and hydathodes of leaves, DAB staining was enhanced in the overall tissues of both cotyledons and leaves of vps29‐3 mutants (Figure S1a). We also performed ROS quantification in the roots of WT and vps29‐3 mutants with ROS‐sensitive fluorescent dye 2′,7′‐dichlorofluorescein diacetate (H₂DCFDA), and the results showed that vps29‐3 mutants accumulated more ROS in their root differentiation zones than WT plants (Figure S1b,c).

Finally, we assessed the transcript levels of PATHOGEN‐RELATED (PR) genes in vps29‐3 mutants because PR genes have been reported to be transcribed upon pathogen‐induced SA accumulation and ROS burst, making them good indicators of immune responses (Saleem et al., 2021; Wu et al., 2012). Our analysis detected significantly increased PR1 and PR2 expression levels in vps29‐3 mutants compared to those in the WT (Figure 1c). These data suggest that the loss of AtVPS29 contributes to the upregulation of SA signaling, ROS accumulation, and subsequent PR gene expression.

We further examined phenotypic correlations by measuring root growth after abscisic acid (ABA) treatment. Root length was shorter in vps29‐3 mutants than in WT plants under untreated conditions, whereas it was similar in WT and siz1‐2 mutants under untreated conditions (Figure S2a). The exogenous application of ABA inhibited root growth in the WT, siz1‐2, and vps29‐3 mutants (Figure S2b). However, the growth inhibitory effect of ABA was greater in siz1‐2 and vps29‐3 mutants than in WT plants, with vps29‐3 mutants showing more sensitivity to ABA than siz1‐2 mutants (Figure S2b). These findings suggest that ABA sensitivity is increased by the loss of AtSIZ1 or AtVPS29, and vps29‐3 mutants are much more sensitive to ABA than siz1‐2 mutants.

AtSIZ1 physically interacts with AtVPS29

Based on the severe dwarf phenotype of vps29‐3 mutants, along with upregulated SA signaling and high ABA sensitivity in these mutants, we hypothesized that the sumoylation system regulated by AtSIZ1 could be affected by VPS29, either directly or indirectly. Therefore, we aimed to investigate the role of VPS29 protein in E3 SUMO ligase activity. To determine whether AtVPS29 affects the stability or activity of AtSIZ1, we first examined the interaction between AtVPS29 and AtSIZ1 using a yeast two‐hybrid assay and immunoprecipitation (IP). Full‐length complementary DNAs (cDNAs) of AtVPS29 and AtSIZ1 were cloned into yeast expression vectors, and their interactions were evaluated. The results demonstrated a strong interaction between AtVPS29 and AtSIZ1 (Figure 2a).

Figure 2.

AtSIZ1 interacts with AtVPS29.

(a) AtSIZ1 directly interacts with AtVPS29 in yeast. Full‐length AtSIZ1 and AtVPS29 cDNAs were fused to sequences encoding the Gal4 activation domain (AD) and the Gal4 DNA‐binding domain (BD) in pGAD424 and pGBT8, respectively. The constructs were transformed into yeast strain AH109. Numbers in each circle quadrant in the left panel correspond to yeast cells transformed with a combination of only pGAD424 and pGBT8 vectors or recombinant plasmids. Transformants were plated onto minimal medium −Leu/−Trp or −Leu/−Trp/−His (5 mm 3‐AT) and incubated for 4 days.

(b, c) In vivo interaction of AtVPS29 and AtSIZ1 in plants was examined using double transgenic plants expressing 35S‐AtVPS29‐Myc ₆ and XVE‐AtSIZ1‐HA ₃ . The plants were incubated in liquid medium supplemented with β‐estradiol to induce the expression of AtSIZ1. After a 15‐h incubation, the levels of AtVPS29‐Myc₆ and AtSIZ1‐HA₃ were assessed by western blotting using anti‐Myc or anti‐HA antibodies (b). Immunoprecipitation was performed using an anti‐Myc antibody, followed by detection of AtSIZ1‐HA₃ through western blotting with an anti‐HA antibody (c). AtVPS29‐Myc₆ was also examined by western blotting using an anti‐Myc antibody. IgG, immunoglobulin G.

For the IP assay, double transgenic plants expressing 35S‐AtVPS29‐Myc ₆ and XVE‐AtSIZ1‐HA ₃ were generated. These plants were then treated with estradiol to induce the expression of AtSIZ1. After confirming the expression of AtVPS29‐Myc₆ and AtSIZ1‐HA₃ (Figure 2b), the total proteins were subjected to IP using an anti‐Myc antibody. The results demonstrated the presence of AtSIZ1‐HA₃ following IP (Figure 2c), indicating a strong interaction between AtVPS29 and AtSIZ1. These findings were consistent with the results obtained from the yeast two‐hybrid experiments.

Pull‐down assays were performed to confirm the interaction between AtVPS29 and AtSIZ1 in an in vitro system. The recombinant proteins were overexpressed in Escherichia coli and purified using affinity columns (Figure S3a,b). The results showed that glutathione S‐transferase (GST)‐AtVPS29, but not GST alone, pulled down AtSIZ1 (Figure S3c). Moreover, experiments with deletion mutants revealed that the N‐terminal region of AtSIZ1, which contained the SAP domain (MBP‐AtSIZ1 [D3]), interacted with AtVPS29 (Figure S3d).

AtVPS29 localizes in the nucleus and in the endosomal membrane

Previous studies have reported that VPS29 primarily functions as part of the retromer complex in the endosomal membrane (Jaillais et al., 2007; Zelazny et al., 2013), indicating that it mainly localizes to cytoplasm‐residing endosomal compartments (Hu et al., 2022; Jaillais et al., 2007). Additionally, it is well known that AtSIZ1 mainly localizes in the nucleus (Kim, Jang, & Seo, 2016; Kwak et al., 2024; Miura et al., 2005). Nevertheless, our analysis revealed that AtSIZ1 directly interacted with AtVPS29 (Figure 2; Figure S3). These findings prompted us to examine the subcellular localization of AtVPS29 in detail. First, we examined the direct interaction and subcellular localization of AtSIZ1 and AtVPS29 using a bimolecular fluorescence complementation (BiFC) assay. Nicotiana benthamiana leaves were coinfiltrated with 35S‐YFP(N)‐AtSIZ1 and 35S‐YFP(C)‐AtVPS29 constructs, followed by analysis using a confocal laser‐scanning microscope (CLSM). The results showed that the YFP signal was primarily detected in the nucleus, with a weak signal also observed in the cytoplasm (Figure 3a), indicating that AtSIZ1 directly interacts with AtVPS29 and that AtVPS29 can localize in the nucleus.

Figure 3.

Subcellular distribution analysis of AtVPS29.

(a) Nicotiana benthamiana leaves were coinfiltrated either with 35S‐YFP(N)‐AtSIZ1 and 35S‐YFP(C) or 35S‐YFP(N)‐AtSIZ1 and 35S‐YFP(C)‐AtVPS29. The agroinfiltrated leaves were incubated at 25°C for 2 days, and then YFP signals were detected by CLSM. Scale bars = 20 μm.

(b) Nuclear and cytosolic fractions were isolated from transgenic plants expressing 35S‐AtVPS29‐HA ₃ , and then AtVPS29‐HA₃ was examined by western blot analysis with an anti‐HA antibody. Actin and Histone 3 were used as nucleus‐ and cytosol‐specific loading controls and detected by anti‐Actin and anti‐histone 3 antibodies, respectively. Cyt, cytosol; Nuc, nucleus.

(c) Detection of AtVPS29 protein in transgenic plants expressing AtVPS29‐GFP. GFP signals were detected by CLSM in the roots from 7‐day‐old plants. Scale bars = 50 μm.

Next, we further examined the nuclear localization of AtVPS29. For this purpose, we isolated nuclear and cytosolic fractions from transgenic plants containing the 35S‐AtVPS29‐HA ₃ construct and extracted the total protein from these fractions. AtVPS29‐HA₃ expression was assessed through western blotting using an anti‐HA antibody. The results demonstrated that AtVPS29‐HA₃ was detected in both the nuclear and cytosolic fractions (Figure 3b). Histone 3, a nuclear marker protein, was exclusively detected in the nuclear fraction, whereas actin, a cytosolic marker protein, was detected only in the cytosolic fraction.

Third, we examined the subcellular localization of AtVPS29 in transgenic plants expressing AtVPS29 under the AtVPS29 promoter. For this experiment, we generated transgenic plants containing the AtVPS29 _pro ‐AtVPS29 cDNA‐GFP construct and detected GFP signals in their roots using CLSM. The results showed that GFP signals were clearly detected in both the nucleus and cytoplasm (Figure 3c). These results support the conclusion that AtVPS29 localizes to both the nucleus and cytoplasm. Additionally, these findings indicate that AtVPS29 can function independently in the nucleus, as well as within the retromer complex in the cytoplasm.

The level of AtSIZ1 protein is positively regulated by AtVPS29

Direct interaction between AtVPS29 and AtSIZ1 implies that the concentration of AtSIZ1 is modulated by the amount of AtVPS29 in vivo. To test this hypothesis, we analyzed AtSIZ1 protein levels in WT and vps29‐3 plants. The results revealed that AtSIZ1 levels were lower in the vps29‐3 mutants than in the WT plants (Figure 4a). We also measured AtSIZ1 concentration in transgenic plants expressing the 35S‐AtVPS29‐HA ₃ construct. The results showed that AtSIZ1 concentrations in plants overexpressing AtVPS29 were approximately 4.64‐fold higher than those in the WT plants (Figure 4c). It is possible that AtSIZ1 transcript levels affect the AtSIZ1 protein concentration. Therefore, we examined AtSIZ1 transcript levels in vps29‐3 mutants and AtVPS29‐overexpressing transgenic plants. The results revealed that AtSIZ1 transcript levels were higher in the vps29‐3 mutants than those in the WT plants (Figure 4b). In addition, AtSIZ1 transcript levels in AtVPS29‐overexpressing plants were similar to those in the WT plants (Figure 4d). Meanwhile, AtVPS29 protein levels and AtVPS29 transcript levels of siz1‐2 mutants were comparable to those of the WT plants (Figure S4). We attempted multiple times to generate vps29‐3 siz1‐2 double mutants by crossing to assess epistasis between AtVPS29 and AtSIZ1. However, we encountered challenges in obtaining double mutants in the F2 generation, suggesting the possible lethality of the double mutants. These results indicate that the level of AtSIZ1 protein is positively regulated by AtVPS29.

Figure 4.

The levels of AtSIZ1 protein in vps29‐3 mutants and AtVPS29‐overexpressing plants.

(a) Total proteins were extracted from 15‐day‐old wild‐type (WT) and vps29‐3 plants. Following 11% SDS‐PAGE, the levels of AtSIZ1 protein were examined by western blotting with anti‐AtSIZ1 antibody. The numbers below the blot represent the relative intensity, with the intensity in WT set to 1. Actin was used as a loading control.

(b) Total RNA was isolated from the same samples described in (a). AtSIZ1 transcript levels were examined using real‐time qRT‐PCR with AtSIZ1‐specific primers. Results are expressed as means ± SD (n = 3). Empty circles indicate individual data points. Asterisks (**) indicate statistically significant differences in transcript levels of AtSIZ1 (P < 0.01; Student's t‐test) between WT and vps29‐3 plants.

(c) Total proteins were extracted from 15‐day‐old WT and AtVPS29‐overexpressing plants. Following 11% SDS‐PAGE, AtSIZ1 protein levels were examined by western blotting with anti‐AtSIZ1 or anti‐HA antibodies. The numbers below the blot represent the relative intensity, with the intensity in WT set to 1. Actin was used as a loading control. Asterisks indicate nonspecific bands.

(d) Total RNA was isolated from the same samples described in (c). AtVPS29 and AtSIZ1 transcript levels were examined using real‐time qRT‐PCR with AtVPS29‐ or AtSIZ1‐specific primers. Results are expressed as means ± SD (n = 3). Empty circles indicate individual data points. Asterisks (***) indicate statistically significant differences in AtVPS29 transcript levels (P < 0.001; Student's t‐test) between WT and AtVPS29‐overexpressing plants (left). NS, not significant (right).

We further examined the effect of AtVPS29 on AtSIZ1 stability in roots using a GFP assay system. Transgenic WT and vps29‐3 mutants containing AtSIZ1 _pro ‐AtSIZ1‐GFP were used for the GFP assay, and GFP signals were monitored using confocal laser scanning microscopy. The results showed lower AtSIZ1‐GFP signals in the root elongation, differentiation, and meristematic zones of the vps29‐3 mutant than in those of the WT plants (Figure 5a; Figure S5). AtSIZ1‐GFP abundance in the vps29‐3 mutant was approximately 65.6 ± 13.2% and 19.2 ± 2.7% compared with that of WT plants in the root meristematic and elongating zones, respectively (Figure 5b). In addition, AtSIZ1‐GFP was observed in the root hairs of WT plants but not in those of vps29‐3 mutants (Figure 5c). The GFP assays support the hypothesis that AtSIZ1 stability is positively regulated by AtVPS29 activity after translation.

Figure 5.

Detection of AtSIZ1 protein in vps29‐3 mutants expressing AtSIZ1‐GFP.

(a) Confocal laser scanning microscopy of roots from 7‐day‐old transgenic WT and vps29‐3 seedlings expressing AtSIZ1‐GFP. Two independent lines in each background were used for the GFP assays. Scale bars = 100 μm.

(b) Quantification of AtSIZ1‐GFP fluorescence intensity. Total fluorescence intensities were measured in the meristematic and elongation zones of the roots of transgenic WT and vps29‐3 seedlings expressing AtSIZ1 _pro ‐AtSIZ1‐GFP (n = 6 independent roots, ***P < 0.001; Student's t‐test). Error bars represent means ± SD. Empty circles indicate individual data points.

(c) Confocal images of the root differentiation zone of seedlings described in (a). Scale bars = 50 μm.

SUMO‐conjugates are less accumulated in vps29‐3 mutants

All the results thus far suggest that the reduced abundance of AtSIZ1 leads to decreased sumoylation of target proteins, which in turn causes phenotypic changes in vps29‐3 mutants. Therefore, the levels of SUMO‐conjugates were examined in WT plants and vps29‐3 mutants under heat stress. A previous study showed that SUMO conjugation was lower in siz1 mutants than in WT plants under normal and heat stress conditions (Yoo et al., 2006). Consistent with previous reports, we found that the levels of SUMO‐conjugates were significantly lower in siz1‐2 mutants than in WT under heat treatment (Figure 6; Figure S6). The levels of SUMO‐conjugates were also much lower in vps29‐3 mutants than in the WT plants under heat treatment (Figure 6; Figure S6). Moreover, even before heat treatment, vps29‐3 mutants displayed lower levels of SUMO‐conjugates than the WT plants, which resembled siz1‐2 mutants (Figure S6). These observations indicate that the decreased levels of SUMO‐conjugates in vps29‐3 mutants result from a reduction in AtSIZ1 levels due to the loss of AtVPS29 activity.

Figure 6.

The levels of SUMO‐conjugates in vps29‐3 and siz1‐2 mutants.

WT, vps29‐3, and siz1‐2 plants were exposed to heat stress (39°C) for 15 min. SUMO‐conjugates were detected by western blotting with anti‐AtSUMO1 antibody.

The level of COP1 protein is negatively regulated by AtVPS29

A previous study reported that the level and function of AtSIZ1 were negatively regulated by the E3 ubiquitin ligase activity of COP1 (Kim, Jang, & Seo, 2016). Since AtSIZ1 levels were lower in vps29‐3 mutants compared with those in the WT, we hypothesized that COP1 may be upregulated in vps29‐3 mutants. Thus, COP1 protein levels were examined in WT and vps29‐3 plants. The results showed that more COP1 protein accumulated in the vps29‐3 mutants than in the WT plants (Figure 7a). We also found that COP1 transcript levels were higher in the vps29‐3 mutant than in the WT plants (Figure 7b). To validate the effect of AtVPS29 on COP1 abundance, we conducted additional experiments to examine COP1 protein levels in AtVPS29‐overexpressing plants. The results revealed a significant decrease in COP1 protein levels in AtVPS29‐overexpressing plants compared to those in WT plants (Figure 7c). Despite the higher transcript levels of AtSIZ1 in vps29‐3 mutants than in the WT plants, the protein levels of AtSIZ1 were decreased in the vps29‐3 mutants. COP1 transcript levels were upregulated and COP1 proteins largely accumulated in vps29‐3 mutants. Additionally, overexpression of AtVPS29 resulted in accumulation of AtSIZ1 and depletion of COP1. These findings indicate that AtVPS29 is involved in the regulation of both AtSIZ1 and COP1 gene transcription, as well as the post‐translational control of AtSIZ1 and possibly COP1 proteins.

Figure 7.

The levels of COP1 protein in vps29‐3 mutants and AtVPS29‐overexpressing plants.

(a) Total proteins were extracted from 15‐day‐old WT, vps29‐3, and cop1‐4 plants. Following 11% SDS‐PAGE, the levels of COP1 protein were examined by western blotting with anti‐COP1 antibody. The numbers below the blot represent the relative intensity, with the intensity in WT set to 1. Actin was used as a loading control. nd, not detected.

(b) Total RNA was isolated from the same samples described in (a). COP1 transcript levels were examined using real‐time qRT‐PCR with COP1‐specific primers. Results are expressed as means ± SD (n = 3). Empty circles indicate individual data points. Asterisks (***) indicate statistically significant differences in COP1 transcript levels (P < 0.001; Student's t‐test) between WT and vps29‐3 mutant plants.

(c) Total proteins were extracted from 15‐day‐old wild‐type (WT) and AtVPS29‐HA₃‐overexpressing plants. Following 11% SDS‐PAGE, the levels of COP1 and AtVPS29 proteins were examined by western blotting with anti‐COP1 or anti‐HA antibodies. The numbers below the blot represent the relative intensity, with the intensity in WT set to 1. Actin was used as a loading control.

hVPS29 positively regulates PIAS1 and negatively regulates huCOP1

AtSIZ1 is an SP‐RING E3 SUMO ligase (Catala et al., 2007). Human PIAS proteins (PIAS1, PIAS3, PIASxa/β, and PIASy) (Kahyo et al., 2001; Sachdev et al., 2001; Schmidt & Müller, 2002) and their yeast homologs (Siz1, Siz2, Mms21, and Zip3) (Cheng et al., 2006; Wilkinson & Henley, 2010) also belong to the SP‐RING E3 SUMO ligase family. Therefore, we investigated whether human PIAS1 is affected by human VPS29 (hVPS29). To address this question, hVPS29‐silenced HeLa cells were generated, and PIAS1 and hVPS29 protein levels were examined through western blotting. The hVPS29 protein was not detected in hVPS29‐silenced HeLa cells (Figure 8a,b). Furthermore, PIAS1 protein levels were lower in hVPS29‐silenced HeLa cells than in non‐silenced cells (Figure 8a). These results indicate that hVPS29 positively regulates PIAS1 levels in the human system.

Figure 8.

huCOP1 and PIAS1 levels in hVPS29‐silenced human cells.

(a) Total proteins were isolated from HeLa cells or hVPS29 siRNA‐treated HeLa cells. Following 11% SDS‐PAGE, PIAS1 and hVPS29 protein levels were examined by western blotting with anti‐PIAS1 or anti‐hVPS29 antibodies. The numbers below the blot represent the relative intensity, with the intensity in Ctrl set to 1. β‐Actin was used as a loading control.

(b) Total proteins were isolated from HeLa cells or hVPS29 siRNA‐treated HeLa cells. Following 11% SDS‐PAGE, huCOP1 and hVPS29 protein levels were examined by western blotting with anti‐huCOP1 or anti‐hVPS29 antibodies. The numbers below the blot represent the relative intensity, with the intensity in Ctrl set to 1. β‐Actin was used as a loading control.

Due to the observed high accumulation of COP1 protein in the vps29‐3 mutants and the depletion of PIAS1 protein in hVPS29‐silenced HeLa cells, we further examined the levels of human COP1 (huCOP1) protein in hVPS29‐silenced HeLa cells. As expected, the levels of huCOP1 protein were found to be higher in hVPS29‐silenced HeLa cells than in non‐silenced cells (Figure 8b), indicating that the loss of hVPS29 led to accumulation of huCOP1 in HeLa cells. These data collectively indicate that the loss of hVPS29 causes accumulation of the E3 ubiquitin ligase huCOP1 and depletion of the E3 SUMO ligase PIAS1 in human cells, showing a pattern similar to that observed with COP1 and AtSIZ1 in Arabidopsis.

AtSIZ1 degradation is delayed in the presence of AtVPS29

To determine whether AtSIZ1 degradation in vps29‐3 mutants is dependent on ubiquitination and the 26S proteasome pathway, we analyzed the effect of the proteasome inhibitor MG132 on the levels of AtSIZ1 in both the WT plants and vps29‐3 mutants. The results demonstrated that the levels of AtSIZ1 increased in both the WT plants and vps29‐3 mutants after MG132 treatment (Figure 9a). However, it appeared that AtSIZ1 accumulated slightly more in the vps29‐3 mutants than in WT plants. Furthermore, we performed a protein degradation assay with and without cycloheximide. The results showed that the degradation of AtSIZ1 was slightly faster in the vps29‐3 mutants than in the WT plants (Figure 9b,c). These data indicate that AtSIZ1 degradation occurs via the 26S proteasome pathway after polyubiquitination and is inhibited by AtVPS29.

Figure 9.

Effects of MG132 or cycloheximide on AtSIZ1 levels.

(a) Degradation of AtSIZ1 via the proteasome pathway in vps29‐3 mutants. Fifteen‐day‐old WT and vps29‐3 mutants were treated with 50 μm MG132, and the levels of AtSIZ1 were examined by western blotting using an anti‐AtSIZ1 antibody. The numbers below the blot represent the relative intensity, with the intensity in WT set to 1. Protein levels were normalized to a value of 1.00 for AtSIZ1 levels in the absence (−) of MG132 in both panels. Actin was used as a loading control.

(b) In vivo degradation rate of AtSIZ1. To assess the degradation of AtSIZ1 in vps29‐3 mutants, 15‐day‐old WT and vps29‐3 plants were incubated in liquid MS medium with 100 μM cycloheximide (CHX). At the indicated time points, protein was extracted and analyzed by western blotting using an anti‐AtSIZ1 antibody. Actin was used as a loading control.

(c) AtSIZ1 levels during degradation were also expressed in a bar chart. The relative protein levels of AtSIZ1 were normalized to numerical values based on a value of 1.0 for the protein levels at 0 hr using the data shown in (b). Empty circles indicate individual data points. Asterisks (*) indicate statistically significant differences in protein levels of AtSIZ1 (P < 0.05; Student's t‐test) between WT and vps29‐3 plants at indicated time points. NS, not significant.

AtSIZ1 ubiquitination is inhibited by AtVPS29

The regulation of AtSIZ1 levels by the E3 ligase activity of COP1 (Kim, Jang, & Seo, 2016) and accumulation of AtSIZ1 in AtVPS29‐overexpressing plants (Figure 4c) prompted us to investigate the effects of AtVPS29 on AtSIZ1 ubiquitination by COP1. Double transgenic plants expressing XVE‐AtVPS29‐Myc ₆ and 35S‐AtSIZ1‐HA ₃ were treated with estradiol to induce the expression of AtVPS29‐Myc₆. After confirming the expression of AtVPS29‐Myc₆ (Figure 10a), the total proteins were subjected to IP using an anti‐HA antibody. The results clearly showed the presence of AtSIZ1‐HA₃ in the immunoprecipitated samples (Figure 10b). Subsequently, the immunoprecipitated samples were analyzed by western blotting using an anti‐ubiquitin antibody to detect ubiquitinated AtSIZ1‐HA₃. The results demonstrated a decrease in the level of ubiquitin‐conjugated AtSIZ1‐HA₃ in the presence of AtVPS29‐Myc₆ (Figure 10c), indicating that AtSIZ1 ubiquitination is inhibited in the presence of AtVPS29.

Figure 10.

The inhibitory effect of AtVPS29 on AtSIZ1 ubiquitination in plants.

(a) Double transgenic plants expressing XVE‐AtVPS29‐Myc ₆ and 35S‐AtSIZ1‐HA ₃ were incubated in liquid medium supplemented with β‐estradiol to induce the expression of AtVPS29‐Myc₆ in the presence of 50 μm MG132. After a 15‐h incubation, the levels of AtVPS29‐Myc₆ were assessed by western blotting using an anti‐Myc antibody. Actin was used as a loading control.

(b) Immunoprecipitation was performed using an anti‐HA antibody, followed by detection of AtSIZ1‐HA₃ through western blotting with an anti‐HA antibody.

(c) Ubiquitinated AtSIZ1‐HA₃ was analyzed by western blotting of immunoprecipitated proteins using an anti‐ubiquitin antibody.

(d) A cell free degradation assay was performed using transgenic plants expressing XVE‐COP1‐Myc ₆ . The expression of COP1‐Myc₆ was induced by β‐estradiol treatment. After 15 h of incubation, COP1‐Myc₆ levels were assessed by western blotting using an anti‐Myc antibody (left). Actin served as a loading control. Cell lysates from β‐estradiol‐treated transgenic plants were incubated with the recombinant protein His₆‐AtSIZ1‐Myc₆ in the presence of GST or GST‐AtVPS29 for the indicated times. The levels of His₆‐AtSIZ1‐Myc₆ were assessed by western blotting using an anti‐Myc antibody (Right). RuBisCO stained with Coomassie Brilliant Blue was used as a loading control. IgG, immunoglobulin G.

To confirm that AtVPS29‐driven inhibition of AtSIZ1 ubiquitination and subsequent degradation is dependent on COP1, we conducted a cell‐free degradation assay using the His₆‐AtSIZ1‐Myc₆ recombinant protein and transgenic plant cell lysates expressing XVE‐COP1‐Myc ₆ . Transgenic plants expressing XVE‐COP1‐Myc ₆ were treated with estradiol to induce the expression of COP1‐Myc₆ (Figure 10d, left). His₆‐AtSIZ1‐Myc₆ purified from E. coli was incubated with the induced cell lysates of XVE‐COP1‐Myc ₆ with the addition of GST or GST‐AtVPS29. The results demonstrated that the degradation of His₆‐AtSIZ1‐Myc₆ was suppressed when GST‐AtVPS29 was added (Figure 10d, right). These data, together with our coimmunoprecipitation results (Figure 10a–c), indicate that AtVPS29 inhibits COP1‐mediated ubiquitination and degradation of AtSIZ1.

AtSIZ1 forms a complex with retromer proteins

There are two possible mechanisms by which AtVPS29 protein regulates SIZ/PIAS‐type E3 SUMO ligase levels. The first possibility is that AtVPS29 alone controls SIZ/PIAS‐type E3 SUMO ligase amounts by directly interacting with AtSIZ1. The second possibility is that AtVPS29 modulates SIZ/PIAS‐type E3 SUMO ligase amounts together with AtVPS26 and AtVPS35, because it functions as part of the VPS26‐VPS29‐VPS35 retromer complex (Seaman, 2021). To investigate the effect of AtSIZ1 on the formation of the trimeric retromer complex (AtVPS29‐AtVPS35a‐AtVPS26a), we purified recombinant proteins, including MBP‐AtSIZ1, GST‐AtVPS29, GST‐AtVPS26a‐HA, and GST‐AtVPS35a‐Myc, and then examined whether MBP‐AtSIZ1 could form a tetrameric complex with GST‐AtVPS29, GST‐AtVPS26a‐HA, and GST‐AtVPS35a‐Myc. The results demonstrated clear detection of all three retromer subunits in the pull‐down assay (Figure S7). These findings indicate that AtSIZ1 can form a complex with retromer proteins through its interaction with AtVPS29.

vps29‐3 mutants display a short hypocotyl under darkness

The pronounced accumulation of the COP1 protein in vps29‐3 mutants suggests that vps29‐3 mutants may exhibit phenotypes opposite to those of cop1 mutants. Furthermore, previous studies on the negative effects of AtSIZ1 on photomorphogenesis prompted us to examine photomorphogenesis‐related traits in vps29‐3 mutants (Lin et al., 2016). To investigate the phenotypes of vps29‐3 mutants, we measured the hypocotyl length and cotyledon angle of WT plants and cop1‐4, vps29‐3, and siz1‐2 mutants. First, we measured the hypocotyl length of these plants grown for 6 days. Interestingly, under light conditions, the hypocotyl length of vps29‐3 mutants was similar to that of cop1‐4 and siz1‐2 mutants, which developed shorter hypocotyls compared to the WT plants (Figure S8a,b). Under 6 days of dark conditions, while the hypocotyl lengths of cop1‐4 mutants were significantly diminished and those of siz1‐2 mutants were slightly reduced compared to those of the WT plants, vps29‐3 mutants developed much shorter hypocotyls than WT plants, siz1‐2, and even cop1‐4 mutants. Additionally, the hypocotyl lengths of dark‐grown WT plants, cop1‐4, vps29‐3, and siz1‐2 seedlings exposed to light for 1 day were similar to those of the dark‐grown WT plants, cop1‐4, vps29‐3, and siz1‐2 mutants (Figure S8a,b). Next, we measured the cotyledon angle of 6‐day‐old seedlings grown under light or dark conditions. The cotyledon angle of vps29‐3 mutants was narrower than that of cop1‐4 and siz1‐2 mutants under light conditions (Figure S8c, left). Surprisingly, despite the short hypocotyls of vps29‐3 mutants under dark conditions, the cotyledons were slightly opened in vps29‐3 mutants, whereas they were widely opened in cop1‐4 mutants (Figure S8c, middle). Even after exposure to light for 1 day, the cotyledons of vps29‐3 mutants remained slightly open (Figure S8c, right). However, the cotyledon phenotypes of vps29‐3 mutants were inconsistent, with abnormal shapes and sizes observed on one side, whereas the other side appeared normal (Figure S8d). Previous studies have also reported severe dwarf phenotypes and abnormal cotyledon development in vps29 mutants (Jaillais et al., 2007; Shimada et al., 2006). When the WT plants and cop1‐4, vps29‐3, and siz1‐2 mutants were grown for 22 days, the shoot parts of vps29‐3 mutants were much smaller than those of the others (Figure S8e). These results indicate that despite the high abundance of COP1 protein, vps29‐3 mutants showed significantly shorter hypocotyl lengths than cop1‐4 and siz1‐2 mutants, and vps29‐3 mutants display severe defects in cotyledon shape and overall growth throughout the vegetative stages.

DISCUSSION

The present work highlights the regulatory role of the VPS29 protein in signal transduction through modulating the stability and activity of SIZ/PIAS‐type E3 SUMO ligases. In animal systems, VPS29, as a component of both the retromer and retriever (DSCR3/VPS29/C16orf62) complexes (Baños‐Mateos et al., 2019; McNally et al., 2017; Phillips‐Krawczak et al., 2015; Singla et al., 2019), serves as a scaffold that physically links the retromer and retriever to various effectors. Consequently, VPS29 is involved in regulating the abundance of hundreds of transmembrane proteins at the cell surface and promoting functional plasticity in cargo recycling (McGough et al., 2014; McNally et al., 2017). In plants, VPS29 is also a component of the retromer complex, which has been implicated in retrograde trafficking from endosomes back to the trans‐Golgi network. The vps29‐3 mutant displays defects in the protein storage vacuoles (Shimada et al., 2006). In addition to its function in vacuolar trafficking, VPS29 has been proposed to be involved in establishing cell polarity during plant organogenesis by influencing the subcellular trafficking of PIN‐FORMED (PIN) proteins (Jaillais et al., 2007). However, its functional role in post‐translational modifications, including sumoylation, has not yet been characterized.

Based on the phenotypic and developmental similarities between siz1 and vps29‐3 mutants (Catala et al., 2007; Jaillais et al., 2007), we investigated SA signaling and ROS levels in vps29‐3 mutants, which were upregulated in siz1 mutants. Although vps29‐3 mutants were smaller than siz1 mutants and exhibited more severe growth and developmental defects, we found that the expression of SA signaling‐related genes and defense‐related PR genes was elevated in vps29‐3 mutants (Figure 1a,c). Additionally, ROS accumulation was significantly higher in vps29‐3 mutants (Figure 1b; Figure S1). These results suggest that the SA‐ and ROS‐mediated regulation of growth and development in siz1 mutants is similarly operative in vps29‐3 mutants, which exhibit comparable dwarf phenotypes. Because SA is a phytohormone responsible for multilayered plant defense and immune responses (Fu & Dong, 2013; Métraux et al., 1990; Vlot et al., 2009; Zhang & Li, 2019) and ROS levels interact with SA signaling pathways (Herrera‐Vásquez et al., 2015), our results also indicate that AtVPS29 plays a critical role in SA‐ and ROS‐mediated defense.

Given the phenotypic correlation between siz1 and vps29‐3 mutants, which suggests that AtVPS29 may be involved in AtSIZ1‐mediated signaling, we investigated the functional role of AtVPS29 protein in the AtSIZ1‐mediated sumoylation system. To explore this, we first examined their interaction and identified a direct interaction between AtVPS29 and AtSIZ1 using yeast two‐hybrid, IP, pull‐down, and BiFC assays (Figures 2 and 3a; Figure S3), suggesting that AtVPS29 influences AtSIZ1.

AtSIZ1 primarily localizes in the nucleus (Kim, Jang, & Seo, 2016; Kwak et al., 2024; Mazur et al., 2018; Miura et al., 2005), whereas VPS29 is found in the endosomal membrane (Hu et al., 2022; Jaillais et al., 2007; Zelazny et al., 2013). This raises questions regarding how these two proteins interact. To investigate this, we examined their interaction and subcellular localization using BiFC, western blotting of fractionated organelles, and a GFP assay with transgenic plants expressing AtVPS29‐GFP (Figure 3a–c). We found that AtSIZ1 directly interacts with AtVPS29 in vivo (Figure 3a). Interestingly, we also discovered that AtVPS29 localizes not only to the cytoplasm but also to the nucleus (Figure 3b,c), suggesting that AtVPS29 can function independently in the nucleus as well as in the membrane as part of the retromer complex.

The direct interaction and colocalization of AtSIZ1 and AtVPS29 suggest that AtVPS29 may influence the level or activity of AtSIZ1. Analysis of the effect of AtVPS29 on the concentration of AtSIZ1 showed a decrease in AtSIZ1 levels in the vps29‐3 mutants (Figure 4a) but a significant increase in AtSIZ1 levels in transgenic plants expressing AtVPS29‐HA₃ (Figure 4c). Further examination of the roots of transgenic WT plants and vps29‐3 mutants containing AtSIZ1 _pro ‐AtSIZ1‐GFP revealed a lower AtSIZ1‐GFP signal in the elongation, differentiation, meristematic zones, and root hairs of vps29‐3 mutants than in those of WT plants (Figure 5; Figure S5). Collectively, these results indicate that AtSIZ1 protein levels are positively regulated by AtVPS29.

Previous studies have shown that SIZ1‐dependent sumoylation of target proteins is induced by heat treatment. For example, the levels of SUMO‐conjugates are much lower in siz1 mutants than in WT plants under heat stress conditions (Yoo et al., 2006). Furthermore, sumoylation of target proteins is induced by heat (Cai et al., 2017; Castro et al., 2012; Kurepa et al., 2003; Miller et al., 2013). Moreover, the rice E3 SUMO ligases OsSIZ1 and OsSIZ2 are involved in the sumoylation of target proteins under heat stress (Kurepa et al., 2003), and transgenic Arabidopsis plants overexpressing the tomato E3 SUMO ligase SlSIZ1 also accumulate SUMO‐conjugates in response to heat stress (Zhang et al., 2017). Evaluation of SUMO‐conjugate levels under heat stress revealed a significant decrease in SUMO‐conjugates in vps29‐3 mutants compared to the WT plants, although SUMO‐conjugates accumulated to a larger extent in vps29‐3 mutants than in siz1‐2 mutants (Figure 6; Figure S6). This result indicates that the decreased levels of SUMO‐conjugates in vps29‐3 mutants result from a reduction in AtSIZ1 levels due to the loss of AtVPS29 activity.

AtSIZ1 degradation by the 26S proteasome complex after polyubiquitination by COP1 activity (Kim, Jang, & Seo, 2016), and the low abundance of AtSIZ1 in vps29‐3 mutants (Figure 4a), suggest COP1 accumulation in vps29‐3 mutants. As expected, the results clearly demonstrated a substantially higher level of COP1 in the vps29‐3 mutants than in the WT plants (Figure 7a), along with a significant decrease in COP1 levels in AtVPS29‐overexpressing plants (Figure 7c). Therefore, these results collectively indicate that the amount of COP1 protein is negatively regulated by AtVPS29.

SP‐RING E3 SUMO ligases are commonly present in humans (PIAS1, PIAS3, PIASxα/β, and PIASγ) (Kahyo et al., 2001; Sachdev et al., 2001; Schmidt & Müller, 2002) and yeast (Siz1, Siz2, Mms21, and Zip3) (Cheng et al., 2006; Wilkinson & Henley, 2010). Therefore, we also extended the evaluation to the level of PIAS1 in hVPS29‐silenced HeLa cells to investigate whether VPS29 may affect the stability of E3 SUMO ligases in humans. Clearly, the levels of PIAS1 were decreased in hVPS29‐silenced HeLa cells (Figure 8a), indicating that VPS29 positively regulates SIZ/PIAS‐type E3 SUMO ligase levels in humans, as also observed in Arabidopsis. Further examination of huCOP1 levels showed an increase in hVPS29‐silenced HeLa cells compared with non‐silenced cells (Figure 8b), indicating that the loss of VPS29 leads to the accumulation of COP1 in humans. These results strongly suggest that VPS29 commonly participates in the modulation of the sumoylation system by SIZ/PIAS‐type E3 SUMO ligases via the E3 ubiquitin ligase COP1 in both plant and animal systems, although it has not yet been determined whether huCOP1 directly ubiquitinates PIAS1 for degradation.

A previous study revealed that loss of COP1 and suppression of COP1 activity cause stabilization of AtSIZ1 in vivo, and cop1‐4 mutants accumulate more SUMO‐conjugates under various abiotic stresses, even before stress treatment (Kim, Jang, & Seo, 2016). In our observations, vps29‐3 mutants accumulated more COP1 protein (Figure 7a) and less AtSIZ1 protein (Figure 4a), and the levels of SUMO‐conjugates before and after heat treatment were significantly lower than those in the WT plants (Figure 6; Figure S6). These collectively suggest that COP1‐mediated AtSIZ1 ubiquitination is responsible for the diminished abundance and activity of AtSIZ1 in vps29‐3 mutants. Our examination of the dependency of AtSIZ1 degradation on the ubiquitination pathway and its degradation rate in vps29‐3 mutants revealed an increase in AtSIZ1 levels after MG132 treatment (Figure 9a) and a slightly faster degradation rate in the vps29‐3 mutants than in the WT (Figure 9b,c). Therefore, these results indicate that AtSIZ1 in vps29‐3 mutants is highly degraded by the 26S proteasome complex after polyubiquitination, and its degradation is prevented by AtVPS29, leading to the stabilization of AtSIZ1. Extended examination of ubiquitin‐conjugated AtSIZ1 levels through double transgenic plants expressing XVE‐AtVPS29‐Myc ₆ and 35S‐AtSIZ1‐HA ₃ revealed a decrease in ubiquitin‐conjugated AtSIZ1‐HA₃ levels after the induction of AtVPS29‐Myc₆ expression (Figure 10c), strongly indicating that AtSIZ1 ubiquitination is inhibited in the presence of AtVPS29. Our cell free degradation assay using transgenic plants expressing XVE‐COP1‐Myc ₆ and the recombinant protein His₆‐AtSIZ1‐Myc₆ and GST‐AtVPS29 showed that even after the induction of COP1, the degradation of His₆‐AtSIZ1‐Myc₆ was protected in the presence of GST‐AtVPS29 (Figure 10d). Altogether, we propose that AtSIZ1 degradation through COP1‐mediated polyubiquitination is inhibited by AtVPS29. The exact mechanism by which AtVPS29 blocks the ubiquitination of AtSIZ1 by COP1 remains to be elucidated, but we carefully infer that a direct interaction between AtSIZ1 and AtVPS29 at the SAP domain of AtSIZ1 (Figures 2 and 3a; Figure S3) may interfere with the binding of COP1 and AtSIZ1 or the E3 ubiquitin ligase activity of COP1.

The functions of the VPS29 protein as a component of the retromer complex and the direct interaction between AtVPS29 and AtSIZ1 suggest that AtSIZ1 can form a complex with trimeric retromer proteins. The results showed that AtSIZ1 could indeed form a complex with the retromer proteins through its interaction with AtVPS29 (Figure S7). These results, along with the nuclear localization of AtVPS29 (Figure 2a–c), suggest that AtVPS29, either alone or as part of a trimeric retromer complex with AtVPS35a and AtVPS26a, can regulate the level and function of AtSIZ1.

Previous studies reported that COP1 overexpression results in elongated hypocotyls under various light conditions (Lin et al., 2016; McNellis, von Arnim, & Deng, 1994). COP1 accumulation in vps29‐3 mutants suggested longer hypocotyl phenotypes in vps29‐3 mutants than in WT plants and cop1 mutants under both light and dark conditions. Meanwhile, siz1 mutants displayed shorter hypocotyls than WT plants under both light and dark conditions, implying that AtSIZ1 negatively regulates photomorphogenesis (Figure S8a,b; Hammoudi et al., 2018; Lin et al., 2016). This suggests the possibility that vps29‐3 mutants can show short hypocotyl phenotypes similar to siz1 mutants. Unexpectedly, however, the hypocotyl length of vps29‐3 mutants was comparable to that of cop1‐4 mutants under light conditions and shorter than that of WT plants, siz1‐2 mutants, and even cop1‐4 mutants under dark conditions (Figure S8a,b). This discrepancy may be explained by the fact that AtSIZ1‐mediated sumolyation of COP1 increases the transubiquitination activity of COP1 on ELONGATED HYPOCOTYL 5 (HY5) (Lin et al., 2016). The impaired SUMO‐conjugating activity of vps29‐3 mutants (Figure 6; Figure S6) can decrease the sumoylation of COP1 and the subsequent E3 ubiquitin ligase activity of COP1 on various photomorphogenesis‐related target proteins, such as HY5 and PHYTOCHROME INTERACTING (PIF) proteins. Previous results showing that siz1 mutation largely rescued the long hypocotyl phenotype of COP1‐overexpressing plants support this notion (Lin et al., 2016). Otherwise, multiple growth and developmental defects in vps29‐3 mutant may contribute to a shorter hypocotyl phenotype, such as its inability to build proper PIN trafficking and cell polarity required for development, an impaired GA signaling pathway, and altered lipid and protein sources in its seeds that are required for proper growth at early stages (Durand et al., 2019; Jaillais et al., 2007; Min et al., 2024; Shimada et al., 2006). Additionally, we encountered difficulties in precisely measuring the cotyledon angle in vps29‐3 mutants. Many vps29‐3 mutants exhibited abnormal cotyledon shapes and sizes, whereas some displayed normal cotyledon phenotypes that remained closed under dark conditions. These findings suggest that the short hypocotyl phenotype in vps29‐3 mutants may result from alterations in various signaling pathways, as well as in COP1‐mediated signal transduction, due to the loss of AtVPS29. Previous studies have reported that loss‐of‐function mutants with various metabolic perturbations frequently exhibit etiolated seedlings with short hypocotyls, even without assessing the effects of COP1. For instance, mutants such as icl‐2, mls‐2, pck1‐2, and fugu5‐1, which involve genes encoding ISOCITRATE LYASE (ICL), MALATE SYNTHASE (MLS), PHOSPHOENOLPYRUVATE CARBOXYKINASE1 (PCK1), and the vacuolar type H+‐pyrophosphatase (FUGU5), respectively, display phenotypic characteristics like short hypocotyls and closed cotyledons in the dark (Ferjani et al., 2011; Tabeta et al., 2022; Takahashi et al., 2017). Therefore, we infer carefully that the hypocotyl phenotypes observed in vps29‐3 mutants, along with their smaller size compared to cop1‐4 mutants (Figure S8e), may result not only from a COP1‐mediated signaling defect but also from disorders in other signaling pathways.

Nonetheless, based on previous studies and the results presented here, we propose possible mechanisms by which AtVPS29 modulates the AtSIZ1‐mediated sumoylation system and signaling through COP1 protein. In WT plants, AtVPS29 downregulates COP1 levels, preventing the ubiquitination of AtSIZ1. Consequently, non‐ubiquitinated AtSIZ1 can sumoylate the target proteins (Figure 11, left). In contrast, in vps29‐3 mutants, COP1 accumulates, leading to the ubiquitination of AtSIZ1. Ubiquitinated AtSIZ1 is then degraded by the 26S proteasome complex, thereby inhibiting AtSIZ1‐mediated sumoylation of target proteins and downstream signaling pathways, including SA signaling (Figure 11, right). Direct interactions between AtSIZ1 and AtVPS29, or with retromer complex subunits, suggest that AtVPS29 may regulate the abundance and activity of AtSIZ1, either independently or as part of the retromer complex through unidentified retromer functions.

Figure 11.

Schematic representation of the regulation of AtSIZ1‐mediated sumoylation system by AtVPS29.

AtVPS29 in WT causes a reduction in COP1 levels leading to AtSIZ1 stability. Stabilized AtSIZ1 then sumoylates target proteins (left). Loss of AtVPS29 leads to COP1 accumulation, which induces AtSIZ1 ubiquitination, leading to reduced sumoylation of target proteins (right). AtVPS29 may also regulate AtSIZ1 through unidentified retromer functions. Ub, ubiquitin.

In conclusion, our study demonstrates that the retromer protein VPS29 positively regulates the sumoylation of target proteins through SIZ/PIAS‐type E3 SUMO ligases. Loss of VPS29 resulted in the depletion of the E3 SUMO ligases, AtSIZ1 in Arabidopsis, and PIAS1 in human cells. Conversely, the E3 ubiquitin ligases COP1 and huCOP1 accumulated in Arabidopsis vps29‐3 mutants and hVPS29‐silenced human cells. Overexpression of AtVPS29 led to the accumulation of AtSIZ1 and depletion of COP1 in transgenic Arabidopsis. Additionally, the degradation rate of AtSIZ1 was faster in the vps29‐3 mutants than in the WT, and AtSIZ1 ubiquitination was inhibited in the presence of AtVPS29. Notably, AtVPS29 localized to both the nucleus and cytoplasm, and AtSIZ1 formed a complex with AtVPS29 and trimeric retromer proteins. Moreover, the loss of AtVPS29 resulted in reduced accumulation of SUMO‐conjugates under heat stress, leading to the upregulation of SA signaling and increased ROS accumulation. Overall, our data strongly suggest that VPS29, whether acting independently or as a part of the retromer complex, plays an essential role in regulating signal transduction by modulating the E3 ligase activity of SIZ/PIAS‐type E3 SUMO ligases in both plants and animals.

MATERIALS AND METHODS

Plant materials, growth conditions, and production of transgenic Arabidopsis plants

The Arabidopsis thaliana Columbia‐0 ecotype (WT), a cop1‐4 mutant (McNellis, von Arnim, Araki, et al., 1994), the T‐DNA insertion knockout mutant siz1‐2, and the T‐DNA insertion knockout mutant vps29‐3 were used in this study. The siz1‐2 (Columbia accession, SALK_065397) and vps29‐3 (Columbia accession, SALK_010106) T‐DNA mutant lines were obtained from the Arabidopsis Biological Research Center (ABRC; Columbus, OH, USA). For plants grown on plates, seeds were surface‐sterilized in commercial bleach containing 5% sodium hypochlorite and 0.1% Triton X‐100 solution for 10 min, rinsed five times using sterilized water, and stratified at 4°C for 2 days in the dark. Seeds were sown on agar plates containing Murashige and Skoog (MS) medium, 1% sucrose, and 0.8% agar, buffered to pH 5.7. For plants grown in soil, the seeds were directly sown in sterile vermiculite. Unless otherwise specified, all plants, including seedlings, were grown at 22°C under a 16 h light/8 h dark cycle in a growth chamber.

The transgenic Arabidopsis plants used in this study were produced as follows. To produce AtVPS29‐overexpressing plants, the corresponding full‐length cDNA was amplified by PCR using a forward primer and a reverse primer tagged with HA₃ and was inserted into the plant expression vector pBA002 (Kost et al., 1998). The recombinant plasmid 35S‐AtVPS29‐HA ₃ was transformed into the Agrobacterium strain GV3101 (GoldBio, St. Louis, MO, USA) via the heat shock method, following the manufacturer's instructions. Using this transformed Agrobacterium, a recombinant plasmid was introduced into WT Arabidopsis via floral dipping (Clough & Bent, 1998). Agar plates containing the antibiotic phosphinotricin (Basta) (Duchefa Biochemie, Haarlem, The Netherlands) were used for selection.

To generate transgenic plants expressing AtVPS29‐GFP, full‐length AtVPS29 cDNA was cloned into the entry vector pENTR3C (Invitrogen, Carlsbad, CA, USA). The promoter regions of AtVPS29 (−3500 bp from the ATG) were amplified by PCR using gene‐specific primers and fused to the entry clone containing AtVPS29 cDNA by treatment with appropriate restriction enzymes and ligation with T4 DNA ligase (New England Biolabs, Ipswich, MA, USA). The recombinant DNA construct AtVPS29 _pro ‐AtVPS29 was introduced into the pMDC107 vector using LR clonase (Invitrogen). The resulting AtVPS29 _pro ‐AtVPS29‐GFP construct was transformed into Agrobacterium strain GV3101 using the heat shock method following the manufacturer's instructions. Using this transformed Agrobacterium, the recombinant plasmid was introduced into WT plants via floral dipping. Agar plates containing the antibiotic hygromycin (Duchefa Biochemie) were used for selection.

To generate transgenic plants expressing AtSIZ1‐GFP, the promoter region of AtSIZ1 (−3535 bp from the ATG) was cloned into the pENTR/D‐TOPO entry vector (Invitrogen). The full‐length AtSIZ1 cDNA was amplified and inserted into an entry clone containing the AtSIZ1 promoter. The recombinant DNA construct AtSIZ1 _pro ‐AtSIZ1 was introduced into the pMDC107 vector using LR clonase (Invitrogen). Agrobacterium transformation, plant transformation, and transgenic plant selection were performed as described above. To produce vps29‐3 mutants expressing AtSIZ1 _pro ‐AtSIZ1‐GFP, transgenic plants stably expressing AtSIZ1 _pro ‐AtSIZ1‐GFP were crossed with vps29‐3 mutants. Three independent lines of the WT plants and vps29‐3 mutants expressing AtSIZ1 _pro ‐AtSIZ1‐GFP were obtained, and the results from two of these lines are presented.

To investigate the interaction between AtSIZ1 and AtVPS29, double transgenic plants were generated. The full‐length cDNA of AtVPS29 was amplified by PCR using a forward primer and a reverse primer tagged with Myc₆. The amplified product was then inserted into the plant expression vector pBA002. Similarly, the full‐length cDNA encoding AtSIZ1 was amplified by PCR using a forward primer and a reverse primer tagged with HA₃, and inserted into the plant expression vector pER8. The resulting recombinant plasmids, 35S‐AtVPS29‐Myc ₆ and XVE‐AtSIZ1‐HA ₃ , were individually transformed into Agrobacterium strain GV3101 using the heat shock method following the manufacturer's instructions. The two Agrobacterium were mixed, and this mixture was used to co‐introduce recombinant plasmids into Arabidopsis plants using floral dipping. Agar plates containing both antibiotics hygromycin and phosphinotricin (Basta) were used for selection.

To investigate the effect of AtVPS29 on the ubiquitination of AtSIZ1, another type of double transgenic plants was generated. The full‐length cDNA of AtVPS29 was amplified by PCR using a forward primer and a reverse primer tagged with Myc₆. The amplified product was then inserted into the plant expression vector pER8. Similarly, full‐length cDNA encoding AtSIZ1 was amplified by PCR using a forward primer and a reverse primer tagged with HA₃ and inserted into the plant expression vector pBA002. Agrobacterium transformation, plant transformation, and transgenic plant selection were performed as described above.

To generate transgenic plants expressing XVE‐COP1‐Myc ₆ for the cell‐free degradation assay, the full‐length cDNA of COP1 was amplified by PCR using a forward primer and a reverse primer tagged with Myc₆. The amplified product was then inserted into the plant expression vector pER8. Agrobacterium transformation, plant transformation, and transgenic plant selection were performed as described above. Primers used for cloning are listed in Table S1.

Construction of recombinant plasmids and purification of recombinant proteins

cDNAs encoding full‐length AtVPS29 were amplified by PCR and inserted into the pGEX4T‐1 vector (Amersham Biosciences, Buckinghamshire, UK) to produce the GST‐AtVPS29 construct. cDNAs encoding AtVPS35a or AtVPS26a were amplified by PCR using a forward primer and a reverse primer tagged with Myc or a reverse primer tagged with HA, respectively. The PCR products were then inserted into the pGEX4T‐1 vector to construct GST‐AtVPS35a‐Myc and GST‐AtVPS26a‐HA, respectively. cDNAs encoding either the full‐length or the deletion mutants of AtSIZ1 cDNA were inserted into the pMAL‐c2x vector (New England Biolabs) to construct maltose binding protein (MBP)‐AtSIZ1 and its deletion mutants. MBP‐AtSIZ1 (D1), MBP‐AtSIZ1 (D2), and MBP‐AtSIZ1 (D3) contained amino acids 90–470, 300–470, and 1–100 of AtSIZ1, respectively.

To generate the recombinant protein His₆‐AtSIZ1‐Myc₆, full‐length cDNA of AtSIZ1 was amplified by PCR using a forward primer and a reverse primer tagged with Myc₆. The PCR products were inserted into the pET‐28a(+) vector to construct His₆‐AtSIZ1‐Myc₆. All constructs were verified by DNA sequencing to ensure that no mutations were introduced. The primers used for cloning are listed in Table S1.

All constructs were transformed into E. coli BL21/DE3 (pLysS) cells. The recombinant proteins were expressed in this E. coli strain using isopropyl‐β‐d‐thiogalactoside (IPTG) treatment and purified according to the manufacturer's instructions. Briefly, for the purification of GST and GST‐fused proteins, bacteria were lysed in phosphate‐buffered saline (PBS) buffer (pH 7.5) containing 1% Triton X‐100, 2 mm phenylmethylsulfonyl fluoride (PMSF), and a proteinase inhibitor cocktail (Roche, Basel, Switzerland). The lysate was then purified using a glutathione resin (Pharmacia, Piscataway, NJ, USA) to obtain GST, GST‐AtVPS29, GST‐AtVPS35a‐Myc, and GST‐AtVPS26a‐HA. For purification of MBP and MBP‐fused proteins, bacteria were lysed in 20 mm Tris–HCl (pH 7.4), 200 mm NaCl, 1 mm ethylenediamine tetraacetic acid (EDTA), 1% Triton X‐100, and 2 mm PMSF, along with a proteinase inhibitor cocktail. Subsequently, the lysate was purified using amylose resins (New England Biolabs) to obtain MBP, MBP‐AtSIZ1 (F), MBP‐AtSIZ1 (D1), MBP‐AtSIZ1 (D2), and MBP‐AtSIZ1 (D3). For the purification of His₆‐tagged proteins, bacteria were lysed in 50 mm NaH₂PO₄ (pH 8.0), 300 mm NaCl, 1% Triton X‐100, 1 mm imidazole, 5 mm DTT, 2 mm PMSF, and a proteinase inhibitor cocktail, and purified on Ni²⁺‐nitrilotriacetate (Ni²⁺‐NTA) resin (Qiagen, Hilden, Germany) to obtain His₆‐AtSIZ1‐Myc₆. Protein concentrations were determined using a Bradford assay (Bio‐Rad, Hercules, CA, USA).

Detection of ROS in Arabidopsis

To analyze the levels of H₂O₂ in the WT, vps29‐3, and siz1‐2 plants, a histochemical staining method using DAB (Sigma‐Aldrich, St. Louis, MO, USA) was adapted (Castro et al., 2022). Briefly, 15‐day‐old WT, vps29‐3, and siz1‐2 plants were pre‐incubated with 10 mm Na₂HPO₄ for 1 day. The plants were vacuum‐infiltrated and then incubated with DAB staining buffer (10 mm Na₂HPO₄ containing 1 mg ml⁻¹ DAB) overnight at room temperature. After staining, the plants were bleached using 96% (v/v) ethanol at 70°C and then photographed.

ROS quantification of root differentiation zones was conducted with ROS‐sensitive fluorescent dye H₂DCFDA (Sigma‐Aldrich). WT and vps29‐3 plants were incubated in liquid MS medium for 1 day. The plants were incubated with 10 μm H₂DCFDA for 20 min at 4°C. After washing with liquid MS, fluorescent signals were observed using a 40× objective with a Leica SP8 X Confocal Microscope (Leica, Wetzlar, Germany). The excitation and emission windows were as follows: excitation at 488 nm and detection at 510–530 nm. Quantification of the GFP signals was performed using ImageJ software (https://imagej.net/software/fiji/).

In vitro binding assay

To examine the in vitro binding of GST‐AtVPS29 to MBP‐AtSIZ1, 2 μg of full‐length MBP‐AtSIZ1 prey and 2 μg of full‐length GST‐AtVPS29 bait were added to 1 ml binding buffer (50 mM Tris–HCl [pH 7.5], 100 mm NaCl, 1% Triton X‐100, 0.2% glycerol, 0.5 mm β‐mercaptoethanol). After incubation at 25°C for 2 h, the reaction mixtures were incubated with a glutathione resin for 2 h before washing six times with buffer (50 mm Tris–HCl [pH 7.5], 100 mm NaCl, 1% Triton X‐100). The absorbed proteins were separated by 11% sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis (PAGE) and detected by western blotting using an anti‐MBP antibody (Santa Cruz Biotechnology, Dallas, TX, USA). To examine the in vitro binding of GST‐AtVPS29 to MBP‐AtSIZ1 deletion mutants, 2 μg of MBP‐AtSIZ1 deletion mutant baits and 2 μg of full‐length GST‐AtVPS29 prey were added to 1 ml binding buffer (50 mm Tris–HCl [pH 7.5], 100 mm NaCl, 1% Triton X‐100, 0.2% glycerol, 0.5 mm β‐mercaptoethanol). After incubation at 25°C for 2 h, the reaction mixtures were incubated with an amylose resin for 2 h before washing six times with buffer (50 mm Tris–HCl [pH 7.5], 100 mm NaCl, 1% Triton X‐100). Absorbed proteins were separated by 11% SDS‐PAGE and detected by western blotting using an anti‐GST antibody (Santa Cruz Biotechnology).

Yeast two‐hybrid assay

The yeast two‐hybrid assay was performed using a GAL4‐based 2‐hybrid system (Clontech, Mountain View, CA, USA). Full‐length AtSIZ1 and AtVPS29 cDNAs were cloned into pGAD424 and pGBT8 (Clontech) to generate the constructs AD‐AtSIZ1 and BD‐AtVPS29. The constructs were transformed into yeast strain AH109 using the lithium acetate method. Yeast cells were grown in a minimal medium (−Leu/−Trp). Transformants were plated on a minimal medium (−Leu/−Trp/−His), including 5 mm 3‐AT, to test the interactions between AtSIZ1 and AtVPS29. The primers used for cloning are listed in Table S1.

BiFC assay

To generate constructs for the BiFC assay, AtSIZ1 and AtVPS29 cDNAs were cloned into pENTR/D‐TOPO and pENTR3C entry vectors (Invitrogen), respectively. Next, AtSIZ1 and AtVPS29 cDNAs were transferred from their respective entry clones to the gateway vector pSAT4‐DEST‐n(174)EYFP‐C1 (ABRC stock number CD3‐1089) or pSAT5‐DEST‐c(175‐end)EYFP‐C1(B) (ABRC stock number CD3‐1097), which contained N‐terminal 174 aa of EYFP (EYFP^N) or C‐terminal 64 aa of EYFP (YFP^C), respectively. The fusion constructs encoding the EYFP(N)‐AtSIZ1 and EYFP(C)‐AtVPS29 proteins were introduced into A. tumefaciens, and the transformed cells were coinjected into N. benthamiana leaves, as described above. YFP signals were detected in the infiltrated leaves using a 40× objective with a Leica SP8 X Confocal Microscope (Leica). The excitation and emission windows were as follows: YFP channel, excitation at 513 nm and detection at 530–550 nm. The primers used for cloning are listed in Table S1.

Total protein extraction and immunoblot

Whole plant samples were ground with liquid nitrogen and homogenized in a protein extraction buffer containing 50 mm Tris–HCl (pH 7.4), 150 mm NaCl, 1% Triton X‐100, and 1 × protease inhibitor cocktail (Roche). After incubation at 4°C for 30 min, samples were centrifuged at 13 000  g for 15 min at 4°C, and the resulting supernatants were boiled in 2 × sample buffer containing 2% SDS at 95°C for 5 min. Protein samples were separated by SDS‐PAGE, transferred to polyvinylidene difluoride (PVDF) membranes, and then immunoblotted with the indicated antibodies. Transferred PVDF membranes were incubated with blocking buffer (1 × Tris‐buffered saline with Tween 20 [TBST], 5% fat‐free skim milk) containing indicated antibodies at 4°C for 16 h.

Separation of nuclear and cytosolic fractions

Three‐week‐old, light‐grown transgenic plants overexpressing AtVPS29‐HA₃ were harvested and ground in liquid nitrogen using a mortar and pestle in the presence of 10 ml grinding buffer (0.3 m sucrose, 40 mm Tris [pH 7.5], 5 mm MgCl₂, 1 mm PMSF, and a protease inhibitor cocktail). The homogenate was filtered through four layers of cheesecloth and then centrifuged at 1500  g for 10 min at 4°C to pellet the nuclei. The supernatant was carefully separated from the pellet and transferred to new tubes. This supernatant fraction was subjected to centrifugation at 12 000  g for 20 min at 4°C, and the resulting supernatant was collected.

To detect the AtVPS29‐HA₃ protein, total protein was extracted from the nuclear and cytosolic fractions and subsequently analyzed by western blotting using an anti‐HA antibody. Similarly, histone 3 and actin were identified through immunoblotting using anti‐histone 3 (Abcam, Cambridge, UK) and anti‐actin (Abcam) antibodies, respectively.

Examination of AtSIZ1, COP1, and AtVPS29 protein levels

Total protein was extracted from WT and vps29‐3 plants grown for 15 days on MS media to examine the relative levels of AtSIZ1 and COP1 proteins in WT and vps29‐3 plants. Following 11% SDS‐PAGE, AtSIZ1 and COP1 protein levels were examined by western blotting using anti‐AtSIZ1 or anti‐COP1 antibodies. To examine AtVPS29, AtSIZ1, and COP1 protein levels in transgenic plants harboring 35S‐AtVPS29‐HA ₃ , total proteins were extracted from WT and AtVPS29‐HA ₃ ‐overexpressing plants grown for 15 days on MS media. Following 11% SDS‐PAGE, AtVPS29‐HA₃, AtSIZ1, and COP1 levels were examined by western blotting using an anti‐HA (Santa Cruz Biotechnology), anti‐AtSIZ1, and anti‐COP1 antibodies, respectively. To examine AtVPS29 protein levels in WT, vps29‐3, and siz1‐2 plants, total proteins were extracted from WT, vps29‐3, and siz1‐2 plants. Following 15% SDS‐PAGE, AtVPS29 levels were examined by western blotting using an anti‐AtVPS29 antibody. Anti‐AtSIZ1 and anti‐COP1 antibodies were kindly provided by Dr. Nam‐Hai Chua of Rockefeller University, USA. Anti‐AtVPS29 antibody was generated as previously described (Min et al., 2024). An anti‐actin antibody was used to detect the loading control actin. Quantification of the relative band intensities was performed using ImageJ software (https://imagej.net/software/fiji/).

Estimation of SUMO‐conjugates

WT, vps29‐3, and siz1‐2 mutants grown on MS medium for 15 days were treated with high temperature (39°C) for 15 min, and then total proteins were extracted from heat‐treated plants. Following 11% SDS‐PAGE, the levels of SUMO‐conjugates were examined by western blotting using an anti‐AtSUMO1 antibody (Abcam). An anti‐actin antibody was used to detect the loading control actin. Experiments were conducted more than 10 times with different biological replicates, and similar results were obtained.

Examination of transcript levels

Plants were grown on plates containing MS medium for 15 days. Total RNA was extracted from WT, vps29‐3, siz1‐2 mutants, and AtVPS29‐overexpressing plants using the RNeasy Plant Mini Kit (Qiagen) following the manufacturer's instructions; quantified; and divided into equal amounts. First‐strand cDNA was synthesized from 5 μg total RNA using an iScript cDNA Synthesis Kit (Bio‐Rad). Equal volumes of cDNA were amplified by real‐time qRT‐PCR (MyiQ; Bio‐Rad) according to the manufacturer's protocol. The specific primers and template cDNA were combined with 25 μl of iQ SYBR Green Super Mix (Bio‐Rad), and the reactions were performed under the following thermal conditions: 50°C for 2 min, 95°C for 10 min, and 40 cycles of 95°C for 15 sec and 60°C for 1 min. The C _T values obtained for the target genes were normalized to the C _T value for ACTIN2, and the data were analyzed using the iCycler IQ software (Bio‐Rad). The PCR primers were designed using Primer3 (http://frodo.wi.mit.edu/cgi‐bin/primer3/primer3.cgi). The amplified PCR products were cloned into the pGEM T‐Easy vector (Promega, Madison, WI, USA), and their sequences were verified by sequencing using an ABI 3730xl DNA Analyzer (Applied Biosystems, Waltham, MA, USA). The primers used for qRT‐PCR are listed in Table S1.

Detection of AtSIZ1‐GFP and AtVPS29‐GFP in roots

Transgenic WT and vps29‐3 plants expressing AtSIZ1 _pro ‐AtSIZ1‐GFP or AtVPS29 _pro ‐AtVPS29‐GFP were grown for 7 days on agar media, and GFP signals were observed using a 40x objective with a Leica SP8 X Confocal Microscope (Leica). The excitation and emission windows were as follows: GFP channel, excitation at 488 nm and detection at 500–550 nm. Quantification of the GFP signals was performed using ImageJ software (https://imagej.net/software/fiji/).

Mammalian cell culture and siRNA‐mediated gene silencing

HeLa cells were purchased from American Type Culture Collection (Manassas, VA, USA). HeLa cells were maintained in Dulbecco's modified Eagle's medium (Invitrogen) supplemented with 10% fetal bovine serum (Gibco, Waltham, MA, USA) and 1% antibiotic/antimycotic (Gibco) in 5% CO₂ in a 37°C incubator. HeLa cells were transfected with a pool of three siRNAs targeting the hVPS29 coding region (5′‐AUGAUGUGAAAGUAGAACGAAUCGA‐3′, 5′‐AUUGUGAGAGGAGACUUCGAUGAGA‐3′, and 5′‐CGAAUACAAAAAACCUUAAAGCCAG‐3′) using Lipofectamine RNAiMAX (Invitrogen) according to the manufacturer's instructions. Cells were transfected with siRNA‐targeting VPS29 or control siRNA (Thermo Fisher Scientific, Waltham, MA, USA) at a final concentration of 20 nm. huCOP1‐silenced HeLa cells were generated with a pool of three siRNAs targeting the COP1 coding region (5′‐AUGAUGUGAAAGUAGAACGAAUCGA‐3′, 5′‐AUUGUGAGAGGAGACUUCGAUGAGA‐3′, and 5′‐CGAAUACAAAAAACCUUAAAGCCAG‐3′) as described above. Protein depletion efficiency was determined by western blotting. The experiments were repeated five times, and one of the representative samples was used for further experiments.

Detection of hVPS29, PIAS1 and huCOP1 in HeLa cells

HeLa cells transfected with siRNAs were treated with 1 μm mitomycin C (Sigma‐Aldrich, USA) for 24 h to induce genome instability. Harvested cells were lysed with MCLB buffer (50 mm Tris, 150 mm NaCl, and 0.5% NP‐40) supplemented with protease inhibitors (Roche) and sonicated for 10 sec. After centrifugation, cleared lysates were separated by SDS‐PAGE, transferred to PVDF membrane (Bio‐Rad), and incubated with the following antibodies: anti‐PIAS1 (Abcam), anti‐hVPS29 (Santa Cruz Biotechnology), anti‐huCOP1 (Bethyl Laboratories, Montgomery, TX, USA), and anti‐β‐actin (Abfrontier, Seoul, South Korea).

In vivo interaction assay

Fifteen‐day‐old light‐grown double transgenic plants of 35S‐AtVPS29‐Myc ₆ and XVE‐AtSIZ1‐HA ₃ were incubated in liquid MS medium with β‐estradiol to induce the expression of AtSIZ1. After incubation for 15 h, total protein was extracted from each sample, and the presence of AtVPS29‐Myc₆ and AtSIZ1‐HA₃ was detected by western blotting using anti‐Myc (Santa Cruz Biotechnology) and anti‐HA antibodies, respectively. Subsequently, total protein was extracted from each sample and immunoprecipitated using an anti‐Myc antibody in a buffer containing 50 mm Tris‐Cl (pH 8.0), 150 mm NaCl, 10% glycerol, 1% NP‐40, 2 mm EDTA, 1 mm PMSF, and a protease inhibitor cocktail. Immunoprecipitated proteins were then analyzed through western blotting using an anti‐HA antibody to detect AtSIZ1. Anti‐actin antibody was used to detect the loading control actin.

Interaction analysis between AtSIZ1 and trimeric retromer proteins

To investigate the in vitro binding of MBP‐AtSIZ1 to retromer proteins, a mixture containing 2 μg of full‐length MBP‐AtSIZ1 and 2 μg of full‐length GST‐AtVPS29, GST‐AtVPS35a‐Myc, and GST‐AtVPS26a‐HA was prepared in 1 ml binding buffer (50 mm Tris–HCl, pH 7.5, 100 mm NaCl, and 1% Triton X‐100). The reaction mixtures were incubated at 25°C for 2 h and then incubated with an amylose resin for an additional 2 h. Afterward, the resin was washed six times using buffer (50 mm Tris–HCl, pH 7.5, 100 mm NaCl, 1% Triton X‐100). Bound proteins were separated through 11% SDS‐PAGE and detected through western blotting using anti‐GST (Santa Cruz Biotechnology), anti‐Myc, and anti‐HA antibodies.

Effects of MG132 and cycloheximide on AtSIZ1 levels

To investigate the effect of the proteasome pathway on AtSIZ1 levels in vps29‐3 mutants, 15‐day‐old WT and vps29‐3 mutants were treated with 50 μm MG132 (Sigma‐Aldrich) for 16 h. Total proteins were extracted from the samples, and the level of AtSIZ1 was analyzed by western blotting using an anti‐AtSIZ1 antibody. An anti‐actin antibody was used to detect the loading control actin.

To examine protein stability, 15‐day‐old WT and vps29‐3 mutants were cultured in liquid MS medium containing 100 μm cycloheximide (CHX; Sigma‐Aldrich) for 4 h. Protein samples were extracted at the indicated time points and analyzed by western blotting using an anti‐AtSIZ1 antibody. An anti‐actin antibody was used to detect the loading control actin.

Detection of ubiquitinated AtSIZ1 in plants

Fifteen‐day‐old light‐grown double transgenic plants of XVE‐AtVPS29‐Myc ₆ and 35S‐AtSIZ1‐HA ₃ were incubated in liquid MS medium with β‐estradiol to induce the expression of AtVPS29‐Myc₆. After incubation for 15 h, total protein was extracted from each sample, and the presence of AtVPS29‐Myc₆ was detected by western blotting using an anti‐Myc antibody. For immunoprecipitation, total protein was extracted from each sample and immunoprecipitated using an anti‐HA antibody in a buffer containing 50 mm Tris‐Cl (pH 8.0), 150 mm NaCl, 10% glycerol, 1% NP‐40, 2 mm EDTA, 1 mm PMSF, and a protease inhibitor cocktail. To detect AtSIZ1‐HA₃ and ubiquitin‐conjugated AtSIZ1‐HA₃, the immunoprecipitated proteins were then analyzed by western blotting using anti‐HA and anti‐ubiquitin antibodies (Santa Cruz Biotechnology), respectively. Anti‐actin antibody was used to detect the loading control actin.

Cell free degradation assay

Fifteen‐day‐old light‐grown transgenic plants of XVE‐COP1‐Myc ₆ were incubated in liquid MS medium with β‐estradiol to induce the expression of COP1‐Myc₆. After incubation for 15 h, the seedlings were harvested, homogenized in liquid nitrogen, and incubated with degradation buffer containing 25 mm Tris–HCl (pH 7.4), 50 mm NaCl, 10 mm MgCl₂, 0.2% Triton X‐100, 1 mm DTT, and 5 mm ATP. Cell lysates were centrifuged at 13 000  g for 10 min at 4°C, and the resulting supernatants were collected. His₆‐AtSIZ1‐Myc₆, GST, and GST‐AtVPS29 protein samples purified from E. coli were incubated with the cell lysates at 30°C for the indicated times and were analyzed by western blotting using anti‐Myc and anti‐GST antibodies.

Phenotypic analysis

Seeds of the WT, cop1‐4, vps29‐3, and siz1‐2 mutants, which were grown and harvested at the same time, were sterilized with 70% ethanol, sown on plates containing MS medium supplemented with 1% (w/v) sucrose and 0.8% (w/v) phytoagar (pH 5.7), and stratified for 3 days at 4°C. To evaluate the effect of light on hypocotyl and cotyledon bending, seedlings were grown for 6 days on plates at 22°C under constant irradiation with white light. For the etiolated growth experiment, the stratified seeds were exposed to light for 8 h and then covered with aluminum foil to cultivate them in a dark chamber at 22°C for 6 days. To examine the effect of light on dark‐grown seedlings, the plates were transferred to continuous light conditions after 6 days of growth in the dark and further incubated for 1 day. Seedlings were photographed after growth under the indicated conditions. The hypocotyl length and cotyledon angle were analyzed for at least 10 individual seedlings using ImageJ software (https://imagej.net/software/fiji/).

Statistical analysis

All statistical analyses were conducted using R version 4.4.1 (https://www.r‐project.org/). Quantification graphs containing means ± standard deviation (SD) were generated using Microsoft Excel software. Differences in the means of all data were determined by Student's t‐test. All experiments were conducted with at least three biological replicates, with similar results.

ACCESSION NUMBERS

The Arabidopsis Information Resource accession numbers of genes used in this study are ACTIN2 (AT3G18780), AtSIZ1 (AT5G60410), AtVPS29 (AT3G47810), COP1 (AT2G32950), VPS26a (AT5G53530), VPS35a (AT2G17790), ICS1 (AT1G74710), EDS5 (AT4G39030), PAD4 (AT3G52430), EDS1 (AT3G48090), PR1 (AT2G14610), and PR2 (AT3G57260).

AUTHOR CONTRIBUTIONS

HSS designed and supervised the work. WKM, JSK, DHK, S‐IK, SWP, JA, SC, M‐JK, and SJL performed the research. WKM, JSK, DHK, S‐IK, SWP, JA, SC, M‐JK, SJL, JTS, YK, and HSS analyzed the data. HSS wrote the paper. All authors read and approved the final manuscript.

CONFLICT OF INTEREST

The authors declare no competing interests.

Supporting information

Figure S1. Levels of reactive oxygen species (ROS) of vps29‐3 mutants.

Figure S2. vps29‐3 mutants are more sensitive to exogenous ABA.

Figure S3. AtVPS29 interacts with the SAP domain of AtSIZ1.

Figure S4. AtVPS29 protein levels and AtVPS29 transcript levels of siz1‐2 mutants.

Figure S5. Confocal images of the meristematic, elongating, and differentiating zones of WT and vps29‐3 seedling roots expressing AtSIZ1‐GFP.

Figure S6. Levels of SUMO‐conjugates in vps29‐3 and siz1‐2 mutant plants under heat stress.

Figure S7. Interaction of AtSIZ1 with trimeric retromer proteins.

Figure S8. Phenotype analysis of vps29‐3 mutants.

Table S1. List of primers used in this study.

DATA AVAILABILITY STATEMENT

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