Arabidopsis thaliana Ubiquitin-Associated Protein 1 (AtUAP1) Interacts with redundant RING Zinc Finger 1 (AtRZF1) to Negatively Regulate Dehydration Response

Abstract

Ubiquitination, one of the most frequently occurring post-translational modifications, is essential for regulating diverse cellular processes in plants during abiotic stress. The E3 ubiquitin (Ub) ligase Arabidopsis thaliana really interesting new gene (RING) zinc finger 1 (AtRZF1) mutation is known to enhance drought tolerance in A. thaliana seedlings. To further investigate the function of AtRZF1 in osmotic stress, we isolated Ub-associated protein 1 (AtUAP1) which interacts with AtRZF1 using a yeast two-hybrid system. AtUAP1, a Ub-associated motif containing protein, increased the amount of Ub-conjugated AtRZF1. Moreover, AtUAP1 RNA interference lines were more tolerant to osmotic stress than wild type, whereas AtUAP1-overexpressing (OX) transgenic lines showed sensitive responses, including cotyledon greening, water loss, proline accumulation and changes in stress-related genes expression, indicating that AtUAP1 could negatively regulate dehydration-mediated signaling. In addition, AtUAP1-green fluorescent protein fusion protein was observed in the nuclei of root cells of transgenic seedlings. Genetic studies showed that the AtRZF1 mutation could rescue the sensitive phenotype of AtUAP1-OX lines in response to osmotic stress, suggesting that AtRZF1 was epistatic to AtUAP1 in dehydration signaling. Taken together, our findings describe a new component in the AtRZF1 ubiquitination pathway which controls the dehydration response in A. thaliana.

Introduction

Ubiquitination regulates the degradation of many proteins in eukaryote cells by post-translational modification. It affects diverse cellular processes including the cell cycle, immune response, hormone signaling, DNA repair and protein trafficking under biotic and abiotic stress conditions (Pickart 2001, Dreher and Callis 2007, Vierstra 2009, Lyzenga and Stone 2012, Sadanandom et al. 2012). Ubiquitination initiates ubiquitin (Ub) activation via Ub-activating enzyme (E1). Ub is then transferred to Ub-conjugating enzyme (E2), forming a thioester-linked E2–Ub intermediate. The substrate-recruiting Ub ligase (E3) then interacts with E2-Ub, allowing for the transfer of Ub to a target protein. E3 ligase is a key factor that determines substrate specificity by recruiting the target protein (Sadanandom et al. 2012). The genome of Arabidopsis thaliana encodes more than 1,400 different E3 ligases, including over 470 really interesting new gene (RING)-type E3 ligases (Stone et al. 2005). Additionally, several studies have reported that RING-type E3 ligases play important roles in abscisic acid (ABA) signaling and abiotic stress responses (Lyzenga and Stone 2012, Chen et al. 2013). For example, RING-type E3 ligase salt- and drought-induced RING finger 1 (SDIR1) can act as a positive regulator of ABA signaling and abiotic stress response. SDIR1 interacts with SDIR-interacting protein 1 (SDIRIP1) to negatively modulate SDIRIP1 stability through the 26S proteasome pathway (Zhang et al. 2007, 2015). Keep on going (KEG) is a RING-type E3 ligase and a negative regulator of ABA signaling through the proteolytic degradation of ABA-insensitive 5 (ABI5), ABA responsive element-binding factor 1 (ABF1) and ABF3 (Stone et al. 2006, Liu and Stone 2010, Chen et al. 2013). Dehydration responsive element-binding protein 2A (DREB2A) interacting protein 1 (DRIP1) and DRIP2 are two RING-type E3 ligases that can negatively regulate the abundance of drought-responsive transcription factor DREB2A (Qin et al. 2008). Furthermore, a new class of Ub factor (E4 factor) that extends the assembly of multi-Ub chains on E3s or substrates has been discovered (Koegl et al. 1999, Weissman 2001). Among a few yeast E4 proteins, Ub fusion degradation 2 (UFD2) was first identified as a poly-Ub chain assembly E4 factor and has been the best characterized to date (Koegl et al. 1999). Yeast E4 UFD2 possesses a conserved C-terminal U-box (Ub fusion degradation homology) domain that mediates the interaction of UFD2 with Ub-conjugated proteins (Koegl et al. 1999).

Several Ub-interacting domains, including Ub-associated (UBA) domain, coupling of Ub conjugation to endoplasmic reticulum degradation (CUE) domain and Ub-interacting motif, have been discovered, offering new opportunities to investigate the molecular mechanisms of Ub recognition (Ponting 2000, Hofmann and Falquet 2001). Previous studies have shown that proteins containing the UBA domain can directly bind to mono-Ub and/or poly-Ub chains, thereby regulating the target substrates degradation through the 26S proteasome (Bertolaet et al. 2001, Chen et al. 2001). The UBA domain is a common motif present in diverse proteins involved in DNA repair, proteolysis, cell cycle progression and stress responses (Lambertson et al. 1999, Ortolan et al. 2000, Bertolaet et al. 2001, Chen et al. 2001). The best-known UBA domain–containing protein is radiation sensitive 23 (RAD23) discovered in yeast (Saccharomyces cerevisiae) through its role in DNA damage repair (Lambertson et al. 1999). RAD23 contains a Ub-like domain in the N-terminus for binding to the 26S proteasome and a UBA domain in the C-terminal region for binding to tetraubiquitin non-covalently (Schauber et al. 1998, Wilkinson et al. 2001). Although many UBA domain–containing proteins in human and yeast have been studied (Bertolaet et al. 2001, Funakoshi et al. 2002), little is known about UBA domain–containing proteins in plants. It has been reported that a rice RAD23 isoform is involved in ABA signaling (Schultz and Quatrano 1997, Zhang et al. 2005). A rice RAD23 can interact with a rice Viviparous 1 (A. thaliana ABI3 ortholog) to modulate ABA responses in germinating seedlings, and Viviparous 1 is degraded by the Ub/26S proteasome system (UPS) (Schultz and Quatrano 1997, Zhang et al. 2005). There are four RAD23 isoforms in A. thaliana. They play an essential role in cell cycle, morphology and fertility of plants by delivering UPS substrates to the 26S proteasome (Farmer et al. 2010).

Previously, we reported that A. thaliana RING zinc finger 1 (AtRZF1), a RING-type E3 ligase, is involved in drought stress response by regulating proline (Pro) metabolism (Ju et al. 2013). We also demonstrated that atrzf1 is tolerant of drought stress, whereas AtRZF1-overexpressing (OX) plants are more sensitive to drought stress than wild-type (WT) plants, suggesting that AtRZF1 can negatively regulate drought response (Ju et al. 2013). In the present study, a yeast two-hybrid screening was performed to identify AtRZF1-interacting proteins. One such interacting protein was a Ub-associated protein designated as AtUAP1. We found that AtUAP1 has E4 ubiquitination factor activity that is involved in increasing the amount of Ub-conjugated AtRZF1. We also found that AtUAP1 was a negative regulator of the water deficit response. Moreover, genetic studies showed that the AtRZF1 mutation could rescue the sensitive phenotype of AtUAP1-OX plants in response to osmotic stress. These results suggest that E3 Ub ligase AtRZF1 is epistatic to E4 Ub factor AtUAP1 in dehydration signaling.

Results

Identification of AtUAP1 as an AtRZF1-interacting protein

AtRZF1, a RING-type E3 ligase, plays important roles in Pro synthesis–mediated drought stress response (Ju et al. 2013). To further characterize the biochemical functions of AtRZF1, we performed a yeast two-hybrid screening of an A. thaliana complementary DNA (cDNA) library. Candidate AtRZF1-interacting proteins were identified through yeast two-hybrid screening using the X-Gal color reaction (Fig. 1A). Overall, this screening identified 11 candidate AtRZF1-interacting clones. Sequence analyses revealed that these candidate clones were ubiquitination-related proteins, a DNA repair protein, ABA receptor, a Ras-related guanosine triphosphate–binding protein, abiotic stress–responsive proteins, protease-related proteins and unknown proteins. One AtRZF1-interacting protein of interest is At5g14540, which harbors a conserved UBA domain at C-terminus similar to the U-box of yeast UFD2 (Azevedo et al. 2001) (Fig. 1B and Supplementary Fig. S1A). Therefore, At5g14540 was designated as A. thalianaubiquitin-associatedprotein 1 (AtUAP1).

Fig. 1

Interaction of AtRZF1 with AtUAP1 and comparison of UBA aa sequences of AtUAP1 homologs. (A) Yeast two-hybrid constructs comprised fragments of AtRZF1 or AtUAP1 fused to the pBD or pAD in vector, respectively. AtRZF1- or AtUAP1-expressing transgenic yeast cells were used in the X-Gal filter assay. AtRZF1 and AtUAP1 co-expressing transgenic yeast cells conferred a positive blue color reaction on X-Gal filter. pAD, yeast expressing pAD vector alone; pBD, yeast expressing pBD vector alone; pAD-AtUAP1, yeast expressing AtUAP1 protein and pBD-AtRZF1, yeast expressing AtRZF1 protein. (B) Schematic representation of C-terminal UBA domain of AtUAP1. The primary structure contains a PR (235–458) and UBA (482–526). Comparison of UBA deduced aa sequences of AtUAP1 with corresponding domain from different plants. aa sequences of AtUAP1 (At5g14540), AtUAP2 (At3g01560), Z. mays UAP1 (ZmUAP1; Zm23681) and O. sativa UAP1 (OsUAP1; Os01g016990) are shown. Identical or similar aa sequences are shaded in black or gray. (C) GST pull-down assay showing the interaction between AtRZF1 and AtUAP1. Left panel, the recombinant His-AtUAP1 was pulled down by GST-AtRZF1 but not by GST control protein alone. His-AtUAP1 fusion protein was detected with an anti-His antibody. Ten percentage of the amount used in the binding reaction is shown. Molecular weights (kDa) of protein standards are shown on the left. Arrow, His-AtUAP1. Right panel, Coomassie Brilliant Blue staining analysis. All experiments were independently repeated four times. Arrow, His-AtUAP1; asterisk, GST-AtRZF1. (D) AtUAP1 interacted with AtRZF1 as shown by BiFC assay. YFP fluorescence from the interaction between AtRZF1-YFPn and AtUAP1-YFPc was observed in the nucleus of the tobacco leaf cells. Co-expression of YFPn and YFPc was used as a negative control. Scale bar, 50 μm.

AtUAP1 cDNA is 1,644 bp in length and encodes a protein of 547 amino acids (aa) with a calculated molecular weight of 60 kDa (Supplementary Fig. S1B). This protein harbored a predicted Pro-rich region (PR) in the central part and a UBA domain in the C-terminal region as revealed by motif scan (http://myhits.isb-sib.ch) (Fig. 1B). AtUAP2(At3g01560) andZm23681andOs01g016990, a homolog and orthologs of AtUAP1 in A. thaliana, Zea mays and Oryza sativa, respectively, were found in the GenBank database (Supplementary Fig. S1B). Multiple sequence alignment revealed that the highly conserved UBA domains of AtUAP2, Zm23681 and Os01g016990 proteins showed 84.4%, 75.6% and 77.8% sequence identity, respectively, with the corresponding part of AtUAP1 (Fig. 1B and Supplementary Fig. S1B). However, the roles of these conserved UBA domains have not been functionally characterized yet. A phylogenetic tree representing distances among A. thaliana, maize and rice UAPs was built using the clustering software MEGA 7.0 (Supplementary Fig. S1C).

To further confirm the interaction between AtUAP1 and AtRZF1, an in vitro protein binding assay was performed (Fig. 1C). Histidine (His)-AtUAP1 recombinant protein was expressed in Escherichia coli and purified under non-denaturing conditions. Purified His-AtUAP1 recombinant protein was incubated with glutathione-S-transferase (GST) or GST-AtRZF1 recombinant protein. The ability of AtUAP1 to interact with GST-AtRZF1 recombinant protein was tested using glutathione-Sepharose resin and an anti-His antibody. As shown in Fig. 1C, AtUAP1 did bind to GST-AtRZF1, but not to GST control protein alone, indicating that AtUAP1 could interact with AtRZF1. Subsequently, bimolecular fluorescence complementation (BiFC) analysis was performed to test the interaction between AtRZF1 and AtUAP1 in planta. AtRZF1 and AtUAP1 were fused to the N- and C-terminal regions of yellow fluorescent protein (YFP) to form AtRZF1-YFPn and AtUAP1-YFPc, respectively, and the two constructs were co-transfected into tobacco plant leaves. The empty vectors (YFPn and YFPc) were used as negative controls. As shown in Fig. 1D, YFP fluorescent signal was observed in the nucleus of the leaf cell co-transfected by AtRZF1-YFPn and AtUAP1-YFPc, while the negative controls failed to yield any fluorescence. These results suggest that both AtUAP1 and AtRZF1 localize to the nucleus, and AtUAP1 interacts with AtRZF1 in the nucleus of plant cells.

Next, in order to determine which domain of AtUAP1 was responsible for interacting with AtRZF1, we divided AtUAP1 into three regions: N-terminal region (NT, 1–234 aa), central region (PR, 235–458 aa) and C-terminal region (UBA, 459–547 aa) (Supplementary Fig. S2A). These three regions were then examined for possible AtRZF1 interaction. Pull-down experiments showed that neither the PR nor the UBA domain of AtUAP1 was involved in the interaction with AtRZF1, whereas the NT domain was sufficient for the interaction with AtRZF1 (Supplementary Fig. S2B). Taken together, our data suggest that AtRZF1 can interact with the NT domain of AtUAP1.

The C-terminal UBA domain of AtUAP1 binds to oligo-Ub chains in vitro

Previous reports have suggested that the UBA domain regulates protein stability by binding or preventing the elongation of multi-Ub chains (Koegl et al. 1999, Chen et al. 2001). Since the C-terminus of AtUAP1 represents a UBA domain (Fig. 1B), we examined whether AtUAP1 could bind to oligo-Ub chains using His-AtUAP1-full length (FL) and His-AtUAP1-NT proteins (Fig. 2A) with various oligo-Ub chains (Ub₂, Ub₃, Ub₄, Ub₅, Ub₆ and Ub₇). After purified His-AtUAP1-FL and His-AtUAP1-NT recombinant proteins were incubated with oligo-Ub chain mixtures, binding complexes were isolated by in vitro pull down with His-resin followed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblot analyses using anti-Ub antibodies (Fig. 2B). After incubating with a mixture of oligo-Ub chains (Ub_2–7), it was found that AtUAP1-FL could bind to Ub₇ chains (red asterisk), while AtUAP1-NT did not bind to any oligo-Ub chains tested (Fig. 2B, left panel). These results indicated that Ub₇ chains could bind to AtUAP1-FL. Next, to verify whether Ub₇ chains could interact with the PR domain or the UBA domain of AtUAP1 (Fig. 2A), western blot analysis was performed using His-AtUAP1-PR and His-AtUAP1-UBA truncated proteins (Fig. 2C). It was found that AtUAP1-UBA protein binds not only to Ub₇ chains, but also to Ub₄ Ub₅ and Ub₆ chains (white asterisks). In contrast, AtUAP1-PR did not interact with oligo-Ub chains as revealed by immunoblot analysis, indicating that only UBA domain of AtUAP1 possessed a binding capacity for oligo-Ub chains of various lengths. Thus, the C-terminal UBA region is essential for oligo-Ub chains binding to AtUAP1 protein. Taken together, these results suggest the possibility that AtUAP1 may function as a Ub chain assembly factor.

Fig. 2

AtUAP1 preferentially binds to Ub₇ chains and oligo-Ub chains binding is dependent on the UBA region at C-terminus of AtUAP1. (A) Constructs of AtUAP1-FL, AtUAP1-NT, AtUAP1-PR and AtUAP1-UBA were used for binding assays with various Ub chains. (B) Ub₇ chains binding to AtUAP1-FL in vitro. Ub moieties were detected by immunoblot analyses with an anti-Ub antibody. Left two lanes, Ub₇ chains were detected with AtUAP1-FL proteins, but not detected with AtUAP1-NT. Red asterisk shows Ub₇ polymers. Middle lane, each Ub chain mixture was loaded. Middle lane shows migration of Ub polymers of various lengths. Right two lanes, each lane was loaded with His-AtUAP1-FL or His-AtUAP1-NT protein, and any bands were not detected by anti-Ub antibodies. Bottom panel, amount of His-AtUAP1-FL and His-AtUAP1-NT used in each pull-down assay as shown by immunoblot analysis with an anti-His antibody. Arrows show His-AtUAP1-FL and His-AtUAP1-NT. (C) Ub_4-7 chains binding to AtUAP1-UBA in vitro. Ub moieties were detected by immunoblot analyses with an anti-Ub antibody. Left two lanes, Ub₄–₇ chains observed with AtUAP1-UBA truncated proteins, but not detected with AtUAP1-PR. White asterisk shows Ub₄, Ub₅, Ub₆ and Ub₇ polymers. Middle lane, each Ub chain mixture was loaded. Middle lane shows migration of Ub polymers of various lengths. Right two lanes, each lane was loaded with His-AtUAP1-PR or His-AtUAP1-UBA protein, and any bands were not detected by anti-Ub antibodies. Bottom panel, amount of His-AtUAP1-PR and His-AtUAP1-UBA used in each pull-down assay as shown by immunoblot analysis with an anti-His antibody. Arrows show His-AtUAP1-PR and His-AtUAP1-UBA.

AtUAP1 is required to enhance poly-Ub chain assembly of AtRZF1

We previously reported that AtRZF1 exhibits Ub E3 ligase activity in vitro (Ju et al. 2013). To investigate whether AtUAP1 could influence the Ub conjugation reaction of the E3 ligase AtRZF1, a recombinant maltose-binding protein (MBP)-AtRZF1 was expressed in E. coli and purified using amylose resin. In the absence of either E1 or E2, ubiquitinated MBP-AtRZF1 was not observed by western blot assay using an anti-Ub antibody (Fig. 3A). In the presence of both E1 and E2, ubiquitinated MBP-AtRZF1 proteins were detected in immunoblot analyses using anti-Ub or anti-MBP antibodies (Fig. 3B, C), consistent with previously published results (Ju et al. 2013). Interestingly, with increasing amounts of AtUAP1-FL protein, the amount of Ub-conjugated AtRZF1 increased (Fig. 3B, C). Additionally, we measured the intensity of ubiquitinated AtRZF1 using the ImageJ program (https://imagej.nih.gov/ij) (Fig. 3D). As shown in Fig. 3, levels of intensity of ubiquitinated AtRZF1 were much higher in the presence of AtUAP1 than that in the absence of AtUAP1. These results suggest that AtUAP1 can enhance the poly-Ub chain assembly of AtRZF1.

Fig. 3

AtUAP1 enhances amount of ubiquitinated AtRZF1 in vitro. (A) Ubiquitination assay of AtRZF1 in the absence of E1 or E2. Omission of E1 or E2 resulted in loss of ubiquitination of AtRZF1. (B and C) Purified MBP-AtRZF1 was incubated at 37°C for 3 h with E1, E2 (UbcH5a), Ub and adenosine triphosphate without or with 1× and 2× His-AtUAP1-FL. Poly-ubiquitination was visualized with anti-Ub (B) or anti-MBP (C) antibody. The addition of AtUAP1-FL markedly enhanced the amount of ubiquitylated AtRZF1 with high molecular weights. Ten percentage input of MBP-AtRZF1 or His-AtUAP1-FL was detected by anti-MBP or anti-His antibodies, respectively (C; bottom panels). (D) Graphs show quantification of relative intensity of ubiquitinated AtRZF1 in the presence of AtUAP1-FL compared to that in the absence of AtUAP1-FL protein. Error bars show standard deviations of three independent experiments. Different letters above the bars indicate statistically significant differences at P < 0.05 by Tukey’s multiple range test.

Expression analysis of AtUAP1 in A. thaliana

To examine the subcellular localization of AtUAP1, we constructed transgenic A. thaliana plants expressing AtUAP1-green fluorescent protein (GFP) under control of the cauliflower mosaic virus 35S promoter. As shown in Supplementary Fig. S3A the green fluorescent signal of AtUAP1-GFP was found in the nuclei of root cells of transgenic seedlings, and the fluorescence of AtUAP1-GFP and 4ʹ,6-diamidino-2-phenylindole (DAPI) that stains nuclei as a control was co-localized in the nucleus. It is considered that fluorescence of both GFP and DAPI outside nuclei is background signal. The result was consistent as shown in Fig. 1D, indicating that AtUAP1 functions in the nucleus.

To obtain insight into the physiological functions of AtUAP1, expression levels of the AtUAP1 gene in the root (Rt), stem (St), leaf (Lf) and flower (Fl) organs were analyzed by quantitative real-time PCR (qPCR). The result showed that AtUAP1 was expressed in four organs with a lesser expression level in the St compared to those in the Rt, Lf and Fl (Supplementary Fig. S3B). In addition, AtUAP1 showed a slightly higher expression in the Lf and Fl than that in the Rt (Supplementary Fig. S3B).

The E3 ligase AtRZF1 is known to be involved in drought stress response (Ju et al. 2013). Thus, we investigated whether the expression level of AtUAP1 might be altered by abiotic stresses. Accumulation of AtUAP1 mRNA in A. thaliana was assessed by qPCR after treatment with mannitol, ABA, salt or hydrogen peroxide (Fig. 4A–D). Results showed that the expression level of AtUAP1 was increased in A. thaliana seedlings by mannitol and ABA, reaching its maximum at 12 h and 6 h after treatment, respectively, followed by a slight decline in its expression (Fig. 4A, B). The transcript level of AtUAP1 reached a peak within 3 h after salt or hydrogen peroxide treatment, after which the level decreased (Fig. 4C, D). Abiotic stress–inducible responsive to ABA 18 (RAB18), responsive to desiccation 29A (RD29A) and AtGSTU5 (for A. thaliana GSTclass tau 5) genes were used as references for mannitol, ABA, salt and oxidative stress treatments (Fig. 4) (Wagner et al. 2002, Ju et al. 2013). These results indicate that AtUAP1 is regulated by dehydration, ABA and oxidative stresses.

Fig. 4

Expression patterns of AtUAP1 in response to dehydration, ABA, salinity and oxidative stress. (A–D) AtUAP1 mRNA levels were determined by qPCR using total RNAs from 14-day-old seedlings treated with 400 mM mannitol (A), 100 μM ABA (B), 150 mM NaCl (C) or 6 mM H₂O₂ (D) for the indicated hours. RAB18, RD29A and AtGSTU5 genes were used as controls for mannitol, ABA, NaCl and H₂O₂ treatment. A. thaliana Act1 was used as an internal control. Error bars indicate standard deviations from three biological replicates with 12 pooled seedlings per genotype per replicate. Different letters above the bars indicate statistically significant differences at P < 0.05 by Tukey’s multiple range test.

Dehydration and ABA responses in AtUAP1 transgenic plants

To examine whether AtUAP1 was associated with the response to osmotic stress and ABA signaling, reverse genetic approaches were taken. We generated AtUAP1-OX and AtUAP1 RNA interference (RNAi) transgenic plants with a WT background. Two independent AtUAP1-OX (OX3–4 and OX4–1) and AtUAP1 RNAi (ri1–4 and ri7–4) lines were selected for further phenotypic characterization. The expression level of AtUAP1 was higher in OX lines while it was lower in RNAi lines than in WT plants as revealed by qPCR (Supplementary Fig. S4A). Seeds of WT and transgenic plants were germinated on half-strength Murashige and Skoog (MS) growth medium (Murashige and Skoog 1962) with or without 400 mM mannitol or 1 μM ABA. The percentage of seedlings with green and fully expanded cotyledons was scored to measure and compare the impact of AtUAP1 transgenic plants on dehydration and ABA responses after 14 days. The percentages of cotyledon greening showed no significant difference between WT and AtUAP1 transgenic plants grown on half-strength MS growth medium (Supplementary Figs. S4B, C). In the presence of 400 mM mannitol, the percentage of cotyledon greening was the most decreased in AtUAP1-OX plants. Only 21.7% of OX3–4 and 25.6% of OX4–1 showed fully expanded green cotyledons. In contrast, 71.1% of ri1–4 and 92.2% of ri7–4 lines showed expanded and green cotyledons. WT plants showed an intermediate phenotype, with 41.1% of seedlings having expanded and green cotyledons (Fig. 5A, B). These results suggest that cotyledon greening in AtUAP1 RNAi lines is tolerant to osmotic stress, whereas AtUAP1-OX transgenic lines are more sensitive to osmotic stress than WT, implying that AtUAP1 could negatively regulate the sensitivity of plants to dehydration stress.

Fig. 5

Phenotypic analysis of AtUAP1 transgenic plants’ response to dehydration stress and ABA. (A and B) Cotyledon greening assays of WT, two independent AtUAP1 RNAi (ri1–4, ri7–4) and two independent AtUAP1-OX (OX3–4 and OX4–1) transgenic lines under dehydration stress (400 mM mannitol). (A) Photograph showing that AtUAP1 RNAi lines have higher cotyledon greening rates than WT and AtUAP1-OX plants. (B) Seedling plants with green cotyledons were counted. Error bars indicate standard deviations from three biological replicates with 60 seedlings per genotype per replicate. Different letters above the bars indicate statistically significant differences at P < 0.05 by Tukey’s multiple range test. (C and D) Cotyledon greening assays under 1 μM ABA condition. (C) Photograph showing that AtUAP1 RNAi lines have higher cotyledon greening rates than WT and AtUAP1-OX plants. (D) The number of seedling plants with green cotyledons was counted. Error bars indicate standard deviations from three biological replicates with 60 seedlings per genotype per replicate. Different letters above the bars indicate statistically significant differences at P < 0.05 by Tukey’s multiple range test. (E) Water loss assay in 14-day-old WT, AtUAP1 RNAi (ri1–4 and ri7–4) and AtUAP1-OX (OX3–4 and OX4–1) transgenic plants. Error bars indicate standard deviations from five biological replicates with four leaves per genotype per replicate. Different letters above the bars indicate statistically significant differences between the plant lines at each time point (P < 0.05, according to Tukey’s multiple range test). (F) Quantification of Pro content in leaves of WT and AtUAP1 transgenic plants. Light-grown 14-day-old plants were grown for 7 days with (normal) or without (drought) watering. Error bars represent standard deviations of three biological replicates. Different letters above the bars indicate statistically significant differences at P < 0.05 by Tukey’s multiple range test.

Next, ABA sensitivity was investigated by measuring the percentage of cotyledon greening. After 14 days of ABA treatment, the percentage of cotyledon greening was the most decreased in AtUAP1-OX lines. Only 26.9% of OX3–4 and 26.8% of OX4–1 showed fully expanded green cotyledons. Conversely, 68.5% of ri1–4 and 77.8% of ri7–4 lines showed expanded and green cotyledons. WT plants showed an intermediate phenotype, with 49.1% of seedlings having expanded and green cotyledons (Fig. 5C, D). It can be concluded that AtUAP1-OX lines were more sensitive to ABA than WT, while AtUAP1 RNAi lines were less sensitive to ABA than WT. These results indicate that AtUAP1 is involved in ABA response. Taken together, these results demonstrate that AtUAP1 is a necessary component for dehydration stress and ABA-modulated early development and growth of A. thaliana.

Analyses of water loss and Pro content of AtUAP1 transgenic plants under drought stress

To further evaluate drought stress response, the water loss of fresh weight (FW) of detached rosette leaves was measured. To assess water loss from leaves, fourth or fifth leaves of similar size and age were detached from WT, two AtUAP1-OX (OX3–4 and OX4–1) and two AtUAP1 RNAi (ri1–4 and ri7–4) transgenic lines and used to measure the water loss of FW as described previously (Ju et al. 2013). After detachment, leaves from AtUAP1 RNAi plants lost less water than those from WT at room-temperature conditions (23^oC, 30% humidity) (Fig. 5E). This result indicates that a reduction of AtUAP1 expression could enhance drought tolerance in plants by reducing the quantity of leaf water loss.

AtRZF1 can act as a negative regulator of Pro synthesis during water deficit stress (Ju et al. 2013). Thus, we analyzed the Pro content in rosette leaves of WT, two AtUAP1-OX (OX3–4 and OX4–1) and two AtUAP1 RNAi transgenic lines (ri1–4 and ri7–4). Under normal growth condition, the Pro contents of all sample seedlings were similar to each other (Fig. 5F). However, Pro content was significantly different between WT and AtUAP1 transgenic plants under drought stress. The Pro contents of the AtUAP1 RNAi lines were increased more after drought treatment than those of WT or AtUAP1-OX lines (Fig. 5F). These results suggest that AtUAP1 negatively regulates Pro production under a drought condition, implying that AtUAP1 might act in a similar pathway to AtRZF1 in response to drought stress.

Expression of osmotic stress–regulated and ABA-metabolic genes in AtUAP1 transgenic plants

To investigate the dehydration response in AtUAP1 transgenic plants at the molecular level, we evaluated expression levels of alternative oxidase 1a (AOX1a), RAB18, delta1-pyrroline-5-carboxylate synthase 1 (P5CS1) and delta 1-pyrroline-5-carboxylate reductase 1 (P5CR1) genes known to be induced by various abiotic stresses (Huang et al. 2008, Ju et al. 2013). Transcript levels of dehydration-inducible genes were increased more in the two AtUAP1 RNAi lines than in WT and AtUAP1-OX transgenic plants following mannitol treatment. In contrast, expression levels of these four genes in AtUAP1-OX lines were lower than those in WT plants under osmotic stress (Fig. 6A–D). These results suggest that AtUAP1 negatively regulates dehydration-related genes. Moreover, it has been reported that ABA-metabolic genes are effectors of osmotic stress signals that regulate plant development and growth (Chan 2012). To investigate whether the osmotic stress–induced phenotypes in AtUAP1 transgenic plants are affected by endogenous ABA level, aldehyde oxidase 3 (AAO3) and cytochrome P450 family 707A1 (CYP707A1) genes (Chan 2012) were chosen in order to analyze the regulation of endogenous ABA metabolism in WT and AtUAP1 transgenic seedlings. Transcript levels of ABA-biosynthetic AAO3 and ABA-catabolic CYP707A1 genes in WT and AtUAP1 transgenic plants exhibited no significant differences under untreated condition (Fig. 6E, F). Under dehydration condition, the transcript level of AAO3 was lower in AtUAP1 RNAi lines than in WT and AtUAP1-OX seedlings, whereas the expression of this gene was slightly different between WT and AtUAP1-OX seedlings (Fig. 6E). In contrast, the transcript level of CYP707A1 was higher in AtUAP1 RNAi lines than in WT and AtUAP1-OX seedlings (Fig. 6F). These results demonstrate that AtUAP1 regulates the expression of ABA-metabolic genes in response to dehydration stress in A. thaliana seedling.

Fig. 6

Expression of dehydration stress–responsive and ABA-metabolic genes in WT and AtUAP1 transgenic plants under osmotic stress condition. (A–D) Dehydration stress–responsive genes AOX1a (A), RAB18 (B), P5CS1 (C) and P5CR1 (D) mRNA levels were determined by qPCR using total RNAs from 14-day-old WT, two individual AtUAP1 RNAi (ri1–4 and ri7–4) and two individual AtUAP1-OX (OX3–4 and OX4–1) seedlings treated without (normal) or with 400 mM mannitol for 6 h. (E–F) ABA-metabolic genes AAO3 (E) and CYP707A1 (F) expression were analyzed by qPCR using total RNAs from 14-day-old WT, two individual AtUAP1 RNAi (ri1–4 and ri7–4) and two individual AtUAP1-OX (OX3–4 and OX4–1) seedlings treated without (normal) or with 400 mM mannitol for 6 h. Act1 mRNA was used as an internal control. Error bars indicate standard deviations from three biological replicates with 12 pooled seedlings per genotype per replicate. Different letters above the bars in (A)–(F) indicate statistically significant differences at P < 0.05 by Tukey’s multiple range test.

AtRZF1 acts downstream of AtUAP1 in response to osmotic stress

To analyze the genetic relationship between AtRZF1 and AtUAP1, we generated AtUAP1-OX plants in the atrzf1 background (AtUAP1 OX2/atrzf1 and AtUAP1 OX17/atrzf1). Expression levels of those transgenic lines were confirmed by reverse transcription PCR (RT-PCR) (Supplementary Fig. S5A). To analyze the percentage of cotyledon greening under dehydration condition, WT, AtUAP1-OX (OX3–4), atrzf1 and AtUAP1 OX/atrzf1 (AtUAP1 OX2/atrzf1 and AtUAP1 OX17/atrzf1) lines were germinated on half-strength MS medium with or without 400 mM mannitol. Under normal growth condition, no difference in cotyledon development was observed among WT, atrzf1 and AtUAP1 transgenic plants (Supplementary Fig. S5B). It has been reported that atrzf1 mutant is tolerant of osmotic stress (Ju et al. 2013). As shown in Fig. 7A, B, while the AtUAP1-overexpression line (OX3–4) was more sensitive to osmotic stress than WT, two individual AtUAP1 OX/atrzf1 double transgenic lines were tolerant of osmotic stress, like the atrzf1 background. Thus, the attenuated osmotic stress response of AtUAP1-OX lines could be rescued by AtRZF1 mutation, indicating that AtRZF1 acts genetically downstream of AtUAP1. Additionally, we generated AtUAP1 RNAi lines with an AtRZF1-OX background (AtUAP1 ri3/AtRZF1 OX and AtUAP1 ri4/AtRZF1 OX). Expression levels of those transgenic lines were confirmed by RT-PCR (Supplementary Fig. S5C). Interestingly, the expression level of AtUAP1 was lower in atrzf1 than that in WT (Supplementary Fig. S5A). Moreover, the transcript level of AtUAP1 was slightly higher in AtRZF1-OX transgenic lines compared to WT (Supplementary Fig. S5C), suggesting that the transcription level of AtUAP1 was positively regulated by AtRZF1. For cotyledon greening measurements under osmotic stress condition, WT, AtUAP1 RNAi (ri7–4), AtRZF1-OX (AtRZF1 OX) and AtUAP1 ri/AtRZF1 OX plants were germinated on half-strength MS medium with or without 400 mM mannitol. Under normal growth condition, no significant difference in cotyledon greening was observed among WT, AtRZF1-OX line and AtUAP1 RNAi transgenic plants during cotyledon development (Supplementary Fig. S5D). We previously reported that AtRZF1-OX lines (AtRZF1 OX) are hypersensitive to osmotic stress (Ju et al. 2013). As shown in Fig. 7C, D, while the AtUAP1 RNAi (ri7–4) line was tolerant of osmotic stress, two individual AtUAP1 ri/AtRZF1 OX double transgenic lines showed more sensitive phenotype to osmotic stress than WT, like the AtRZF1 OX background, implying that the AtRZF1 gene could inhibit the tolerant phenotype of the AtUAP1 RNAi line in response to osmotic stress.

Fig. 7

Genetic interaction between AtRZF1 and AtUAP1 in osmotic stress response. (A and B) Cotyledon greening assays of WT, atrzf1, AtUAP1 OX3–4 and AtUAP1 OX/atrzf1 (AtUAP1 OX2/atrzf1 and AtUAP1 OX17/atrzf1) transgenic lines under osmotic stress condition (400 mM mannitol). (A) Photograph showing that atrzf1 and AtUAP1 OX/atrzf1 lines have higher percentages of cotyledon greening than WT and AtUAP1-OX OX3–4 plants. (B) The number of seedling plants with green cotyledons was counted. Error bars indicate standard deviations from three biological replicates with 60 seedlings per genotype per replicate. Different letters above the bars indicate statistically significant differences at P < 0.05 by Tukey’s multiple range test. (C and D) Cotyledon greening assays of WT, AtRZF1-OX, AtUAP1 RNAi (ri7–4) and AtUAP1 RNAi/AtRZF1-OX (AtUAP1 ri3/AtRZF1 OX, AtUAP1 ri4/AtRZF1 OX) transgenic lines under osmotic stress condition (400 mM mannitol). (C) Photograph showing that the AtUAP1 RNAi line had higher percentages of cotyledon greening than WT, AtRZF1 OX and AtUAP1 RNAi/AtRZF1-OX lines, while AtRZF1 OX and AtUAP1 RNAi/AtRZF1-OX lines had lower percentages of cotyledon greening than WT. (D) The number of seedling plants with green cotyledons was counted. Error bars indicate standard deviations from three biological replicates with 60 seedlings per genotype per replicate. Different letters above the bars indicate statistically significant differences at P < 0.05 by Tukey’s multiple range test.

Additionally, to further investigate whether the AtRZF1 influences the mRNA levels of dehydration-inducible genes in AtUAP1 OX/atrzf1 and AtUAP1 ri/AtRZF1 OX lines under osmotic stress conditions, a second set of tests was conducted by qPCR assay using AOX1a, RAB18, P5CS1 and P5CR1 genes. As shown in Supplementary Fig. S6 while the transcript levels of all these genes in the AtUAP1-OX plant were decreased compared to the WT, the expression levels of all these genes in two individual AtUAP1 OX/atrzf1 double transgenic lines were higher than those in WT after mannitol treatment, which is similar to the transcript levels of the atrzf1 background. In contrast, the transcript levels of all these genes in two individual AtUAP1 ri/AtRZF1 OX double transgenic lines were decreased compared to the WT after mannitol treatment, which is similar to the transcript levels in AtRZF1 OX background, while the expression levels of all these genes in AtUAP1 RNAi line was increased more than in WT. These results were consistent with phenotypic analysis data, suggesting that AtRZF1 is epistatic to AtUAP1 in response to dehydration.

Discussion

Identification of AtUAP1 as a protein with E4 functions

Ubiquitination plays an important role in the perception and signal transduction of various internal hormones and external environmental signals (Dreher and Callis 2007, Qin et al. 2008, Lyzenga and Stone 2012). Previously, we demonstrated that AtRZF1 is an E3 ligase that acts negatively in Pro synthesis (Ju et al. 2013). We also previously found that AtRZF1 is involved in plant response to drought stress (Ju et al. 2013). However, its targets or interacting proteins remain unknown. In this report, a AtUAP1 gene encoding a protein with a conserved UBA motif was identified as an AtRZF1-interacting protein using yeast two-hybrid screening, pull-down assays and in vivo BiFC analysis (Fig. 1). Furthermore, Supplementary Fig. S2 shows that the NT domain of AtUAP1 was necessary for the interaction with AtRZF1.

Although numerous reports have suggested that ubiquitination pathway–related genes either positively or negatively regulate plant abiotic stress response (Qin et al. 2008, Liu and Stone 2010, Zhang et al. 2015), proteins containing UBA motifs and the effects of UBA motifs on the ubiquitination pathway in plant responses to abiotic stress remain to be elucidated. Yeast and human RAD23 is a protein containing a UBA domain that can non-covalently interact with Ub chains and inhibit the assembly of substrate-linked multi-Ub chains (Ortolan et al. 2000), suggesting that the UBA domain controls protein stability by preventing the expansion of nascent multi-Ub chains (Chen et al. 2001). A previous study reported that yeast UFD2 possesses a conserved C-terminal U-box domain, which mediates the interaction of UFD2 with Ub-conjugated proteins. Moreover, UFD2 promotes the formation of the multi-Ub chain assembly as an E4 factor (Koegl et al. 1999, Ferreira et al. 2015), demonstrating that UFD2 has an antagonistic role to RAD23. E4 factors can catalyze poly-Ub chain assembly in conjunction with E1, E2 and E3 enzymes (Koegl et al. 1999, Ortolan et al. 2000). UFD2 can efficiently facilitate poly-ubiquitination of Ub-Protein A, a UFD2 substrate, together with E1, E2 and E3 enzymes, but not with just E1 and E2, suggesting that E3 is required for E4 function on certain substrates (Koegl et al. 1999). In A. thaliana, the E4 factor MUSE3 (for Mutant, snc1-enhancing 3) is involved in the poly-ubiquitination of nonexpressor of pathogenesis-related 1 (NPR1), leading to its proteasomal degradation (Skelly et al. 2019). The current working model for E4 function in yeast, animals and plants suggests that E4 and its substrate complex may be formed to regulate the transfer of Ub from E2 to the substrate or sequentially add Ub to the substrate after E3 ligase first ubiquitinates the target protein (Koegl et al. 1999, Hoppe 2005, Skelly et al. 2019).

As shown in Fig. 2, we demonstrated that AtUAP1 is capable of binding to poly-Ub chains in vitro. AtUAP1-UBA truncated protein could bind not only to Ub₇ chains, but also to Ub₄ Ub₅ and Ub₆ chains, whereas AtUAP1-FL only interacted with Ub₇ chains. We assumed that the PR domain of AtUAP1 may function to block Ub₄ Ub₅ or Ub₆ chain interaction as the NT domain of AtUAP1 is required for AtRZF1 interaction (Supplementary Fig. S2). Previously, Rotem et al. (2008) reported that the Pro-rich domain of apoptosis-stimulating proteins of p53 (ASPP2) functions to control the interactions of ASPP2 with other proteins. These findings suggest the possibility that the PR domain of AtUAP1 may control the binding of Ub oligomer chains to the UBA domain of AtUAP1, whereas the NT domain of AtUAP1 is involved in the protein–protein interaction.

Intriguingly, we found that AtUAP1 could increase the amount of Ub-conjugated AtRZF1 (Fig. 3). Koegl et al. (1999) reported that the efficient multi-ubiquitination needed for proteasomal targeting of a substrate requires an additional E4 UFD2 protein in yeast. Therefore, we consider that the function of AtUAP1 is similar to that of yeast UFD2 in facilitating the formation of multi-Ub chain assembly. The AtRZF1–AtUAP1 complex may be formed to coordinate the transfer of Ub to their target proteins. In addition, we found that AtUAP1 was localized in the nucleus (Fig. 1D and Supplementary Fig. S3A), implying that AtUAP1 functions as an Ub chain assembly factor in the nucleus.

Physiological roles of the AtUAP1–AtRZF1 module in osmotic stress tolerance

The expression level of AtUAP1 was increased by osmotic stress, ABA, salt and oxidative stress (Fig. 4), suggesting that AtUAP1 is involved in abiotic stress responses. As shown in Fig. 5A–D, AtUAP1-OX lines were more sensitive to osmotic stress and ABA treatment than WT during cotyledon development, whereas AtUAP1 RNAi transgenic lines were less sensitive to osmotic stress and ABA treatment than WT. Moreover, Fig. 6E, F suggest more complicated (and/or redundant) regulation of osmotic stress response by AtUAP1 and AtRZF1. These physiological and molecular data showed that AtUAP1 could negatively regulate dehydration and ABA responses by a ubiquitination-mediated pathway.

Pro accumulation is a common physiological response in plants exposed to drought and salt stress (Hare et al. 1999). Pro is considered an important osmolyte that acts as a molecular chaperone to stabilize the structure of proteins. It also acts as a regulator of cellular redox potential and an antioxidant to control free radical levels (Hare et al. 1999). The present study also demonstrated differences in the water loss and Pro content between AtUAP1-OX and AtUAP1 RNAi transgenic lines (Fig. 5E, F). Leaves of AtUAP1 RNAi lines exhibited a significantly higher water content under drought condition than leaves of WT and AtUAP1-OX lines (Fig. 5E). The accumulation of Pro in the leaves of AtUAP1 RNAi lines was greater than that in WT and AtUAP1-OX plants (Fig. 5F), suggesting that AtUAP1 might be a component responsible for regulating leaf drought sensitivity by modulating compatible solutes. It was shown previously that Pro accumulation under water deficit condition is correlated with dehydration stress tolerance and that its concentration can change the expression of genes involved in the abiotic stress response of plants (Hare et al. 1999, Ju et al. 2013).

In the present study, transcript levels of drought-inducible genes including AOX1a, RAB18, P5CS1 and P5CR1 were significantly increased in AtUAP1 RNAi lines compared to WT following osmotic stress treatment (Fig. 6), suggesting that AtUAP1 could participate in the osmotic stress signaling required to regulate the expression of dehydration stress–related genes.

Genetic relationship between AtUAP1 and AtRZF1

To understand the interaction between AtRZF1 and AtUAP1 in osmotic stress genetically, we generated AtUAP1-OX plants in the atrzf1 background and AtUAP1 RNAi in the AtRZF1-OX background (Fig. 7 and Supplementary Fig. S5). Interestingly, AtUAP1 expression was reduced in the atrzf1 mutant but increased in the AtRZF1-OX line compared to WT under normal condition (Supplementary Figs. S5A, C). These results suggest that the expression level of AtUAP1 might be regulated through the ubiquitination pathway in which AtRZF1 biochemically functions. As shown in Fig. 7, atrzf1, AtUAP1 RNAi and AtUAP1 OX/atrzf1 transgenic lines were less sensitive to osmotic stress than WT and AtRZF1- and AtUAP1-OX plants, whereas AtRZF1- and AtUAP1-OX plants were more sensitive than WT. In addition, there was no significant difference in sensitivity to osmotic stress among atrzf1, AtUAP1 RNAi and AtUAP1 OX/atrzf1 lines. Furthermore, AtUAP1 ri/AtRZF1 OX double transgenic lines were more sensitive to osmotic stress than WT, atrzf1 and AtUAP1 RNAi seedlings. As shown in Supplementary Fig. S6 the highly upregulated expression of dehydration-inducible genes in the AtUAP1 RNAi line was reversed by AtRZF1 overexpression. These results suggest that in the situations where AtRZF1 exists in excess, the functions of AtUAP1 diminish, indicating that AtRZF1 acts genetically downstream of AtUAP1 in response to osmotic stress.

In conclusion, nuclear-localized AtUAP1 as an E4 factor is a negative regulator of ABA and dehydration stress response. Genetic studies revealed that AtRZF1 was epistatic to AtUAP1. Moreover, AtUAP1 could biochemically bind to oligo-Ub chains attached to E3 ligase AtRZF1 and enhance the amount of ubiquitinated AtRZF1. Identifying substrates of AtRZF1 may shed more light on the impact of the increased poly-Ub assembly of AtRZF1.

Materials and Methods

Yeast two-hybrid screening

Yeast two-hybrid screening was conducted following the protocol described previously (Xie et al. 1999). Briefly, AtRZF1 was fused to the Galactose-responsive transcription factor (GAL4) DNA-binding domain in GAL4 DNA-binding domain (pBD) vector as bait. For initial screening, A. thaliana GAL4 activation domain/cDNA library as prey was constructed using GAL4 activation domain (pAD) vector (Stratagene, La Jolla, CA, USA). Then, cDNA clones in pAD vector and pBD-AtRZF1 construct were co-transformed into yeast strain Y190 using the lithium method of Gietz and Schiestl (2007). Transformants were then selected on Trp- and Leu-free synthetic dextrose medium supplemented with 10 mM 3-aminotrizole. After 3 days of culturing at 30°C, primary positive colonies were re-plated on selection plates and subjected to X-Gal filter assays as previously described (Xie et al. 1999). Candidate AtUAP1-expressing yeast colonies were lifted off and used as templates for colony PCR. Reactions primers were as follows: Forward (5ʹ-AGGGATGTTTAATACCACTAC-3ʹ) and Reverse (5ʹ-GCACAGTTGAAGTGAACTTGC-3ʹ). Amplicons were inserted into pGEM T-easy vector (Promega, Madison, WI, USA) for DNA sequencing.

Glutathione S-transferase pull-down assay

FL coding region of AtRZF1 cDNA was amplified with EcoRI-F1 and SalI-R1 primer set (Supplementary Table S1). PCR products were digested with restriction enzymes EcoRI and SalI and inserted into pGEX4T-3 vector. FL AtUAP1 cDNA was amplified with EcoRI-F2 and SalI-R2 primer set (Supplementary Table S1). PCR products were digested with restriction enzymes EcoRI and SalI and inserted into pET-28a vector. Then, each resultant plasmid was transformed into E. coli BL21 to induce GST-AtRZF1 or His-AtUAP1-FL proteins. GST pull-down assay was conducted as described previously (Qin et al. 2008). GST and GST-AtRZF1 recombinant proteins were incubated with 40 µl of a 50% slurry of glutathione-Sepharose 4B beads at 4°C for 2 h in binding buffer [50 mM hydroxyethylpiperazine ethane sulfonic acid (HEPES), pH 7.5, 1 mM ethylenediaminetetraacetic acid (EDTA), 150 mM NaCl, 0.1% Tween 20, 0.5 mM dithiothreitol (DTT) and 0.8% glycerol]. After washing with the binding buffer, the His-AtUAP1-FL recombinant protein was added and incubated at 4°C for 2 h. The mixture was washed three times with 1 ml of binding buffer. Washed beads were boiled with SDS sampling buffer at 100°C for 5 min. Then, the eluted proteins were separated by 10% SDS-PAGE and detected by western blot using an anti-His-antibody. Ten percentage input was loaded as a control to indicate the original amount of prey protein.

BiFC assay

BiFC assay was performed as described previously (Zhu et al. 2019). The coding region of AtRZF1 or AtUAP1 was amplified and inserted into 35S-YFPn (pGTQL 1211YN) or 35S-YFPc (pGTQL 1221YC), respectively, to generate AtRZF1-YFPn or AtUAP1-YFPc constructs. The Agrobacterium tumefaciens strain GV3101 that contains AtRZF1-YFPn or AtUAP1-YFPc was co-introduced into the leaves of tobacco (Nicotiana benthamiana). After incubation for 3 days, the leaves were collected for observation under laser-scanning confocal microscope (Carl Zeiss AG, LSM 800, Oberkochen, Germany). The fluorescence images were obtained and processed using the Carl Zeiss ZEN 2012 SP2 software.

Ub chain binding assay

For Ub-binding assays, AtUAP1-NT, AtUAP1-PR and AtUAP1-UBA truncated constructs were amplified by PCR using primer sets harboring EcoRI and SalI sites (Supplementary Table S1). Coding sequences were inserted into pET-28a vector. His-AtUAP1-FL, His-AtUAP1-NT, His-AtUAP1-PR and His-AtUAP1-UBA used as bait proteins were expressed in E. coli and purified using Ni-NTA agarose (Qiagen, Hilden, Germany). These His fusion proteins were diluted to 200 pmol/50 μl using 50% slurry of lysis buffer (50 mM sodium phosphate monobasic, 300 mM sodium chloride and 10 mM imidazole, pH 8.0). Then 50 μl of the His fusion proteins in 450 μl of lysis buffer were mixed with 0.5 μg of Ub or oligo-Ub ranging from Ub₂ to Ub₇ (Enzo Life Sciences, Farmingdale, NY, USA) as prey. Samples were incubated at 4°C for 2 h. Beads were washed three times with 1 ml of lysis buffer, stopped with SDS sample buffer, separated by 15% SDS-PAGE and analyzed by immunoblotting with an anti-Ub antibody. Inputs were loaded as controls to indicate original amounts of prey and bait proteins and non-specific bands.

In vitro ubiquitinylation assay

In vitro ubiquitinylation assay was performed as described previously (Ju et al. 2013) using an Auto-ubiquitinylation kit (Enzo Life Sciences, Farmingdale, NY, USA). The purified FL E3 ligase AtRZF1-fused to MBP and components required for ubiquitinylation reaction were added to each reaction in the absence or presence of His-AtUAP1-FL. After 3 h of incubation at 37°C, reactions were stopped with SDS sample buffer, separated by 7.5% SDS-PAGE and analyzed by immunoblotting with anti-Ub (Enzo Life Sciences, Farmingdale, NY, USA) or anti-MBP (New England BioLabs, Beverly, MA, USA) antibodies.

Subcellular localization of AtUAP1-GFP in transgenic root cells

FL AtUAP1 cDNA was amplified by PCR using primer set (Supplementary Table S1). PCR products were initially cloned into pDONR/ZEO vector and confirmed by sequencing. Resultant plasmids were then directly cloned into a plant expression vector pEarleygate103 (Earley et al. 2006) under the control of the constitutive 35S promoter. The construct was transformed into WT (Columbia-0) using A. tumefaciens strain GV3101 via in planta vacuum infiltration. Phosphinothricin (Duchefa, Haarlem, Netherlands) resistance of T2 generation from these selected lines segregated as a single locus. Resultant T3 homozygous transgenic 35S promoter-AtUAP1-GFP plants were used to analyze the subcellular localization of AtUAP1. The reagent 4ʹ,6-diamidino-2-phenylindole (DAPI) (Sigma, St. Louis, MO, USA) was used to stain nuclei. Root samples were mounted on microscope slides and observed using a FluoView1000 confocal microscope (Olympus, Tokyo, Japan). Confocal images were obtained and processed using FV10-ASW 1.7W computer software (Olympus, Tokyo, Japan).

Total RNA extraction, RT PCR and qPCR analysis

Total RNAs were isolated from frozen samples using the RNeasy Plant Mini kit (Qiagen, Valencia, CA, USA). To remove residual genomic DNA from each preparation, total RNAs were treated with RNase-free DNase I (Qiagen, Valencia, CA, USA) following the manufacturer’s instructions. They were then used for reverse transcription using SuperScript II reverse transcriptase (Invitrogen) according to the supplier’s instructions. RT-PCR was employed to measure expression levels of AtUAP1 or AtRZF1 in transgenic plants. The amount of RNA used in RT-PCR was 300 ng. After 28 PCR cycles for amplification, 20 µl of each RT-PCR product was loaded onto a 1.0% (w/v) agarose gel to visualize the amplified DNA. qPCR was performed using a CFX Connect quantitative PCR apparatus (Bio-Rad, Hercules, CA, USA). iQ SYBR Green Supermix kit (Bio-Rad) was used for qPCR following the manufacturer’s instructions. Each result was analyzed using CFX maestro software (Bio-Rad). Quantitative analyses were performed using the Delta Delta C_T method (Livak and Schmittgen 2001). Each sample was used for independent experiments. Actin 1 (ACT1) was used as an internal control. Primers used for RT-PCR and qPCR are shown in Supplementary Table S1.

Plant materials, growth conditions and stress induction

In the present study, seeds of A. thaliana (Columbia-0) and atrzf1 plants previously described by Ju et al. (2013) were used. A. thaliana seedlings were grown in a growth room under intense light at 22°C with a relative humidity of 60% and a 16-h day length. For ABA or salt stress, 14-day-old A. thaliana seedlings were submerged in a solution containing 100 μM ABA or 150 mM NaCl and sampled at 0, 3, 6 and 12 h. For osmotic stress, 14-day-old A. thaliana seedlings were submerged in a solution containing 400 mM mannitol and sampled at 0, 6, 12 and 36 h. For oxidative stress, 14-day-old A. thaliana seedlings were submerged in a solution containing 6 mM H₂O₂ and sampled at 0, 1, 3 and 6 h. For drought stress, seedlings were grown in pots with normal watering every 4 days. After 14 days, plants were divided into two groups for stress treatments. One group was subjected to drought stress by withholding water for 7 days. The control group was watered normally. In each case, collected seedlings were promptly frozen in liquid nitrogen and stored at −80°C.

Construction and generation of AtUAP1 transgenic lines

To generate AtUAP1 RNAi lines, 200-bp fragment of AtUAP1 cDNA was amplified by PCR using RNAi primer set (Supplementary Table S1). PCR products were initially cloned into pDONR/ZEO vector and confirmed by sequencing. Resultant plasmids were then directly cloned into pK7GWIWG2D(II) RNAi vector fused with the constitutive 35S promoter (Karimi et al. 2002). The resultant construct was transformed into WT or AtRZF1-OX plants using A. tumefaciens strain GV3101 via in planta vacuum infiltration. T₂ transformants (kanamycin resistance) were selected from transgenic lines segregated as a single locus. T₃ homozygous transgenic AtUAP1 RNAi lines (ri1–4 and ri7–4) and AtUAP1 RNAi/AtRZF1-OX lines (AtUAP1 ri3/AtRZF1 OX and AtUAP1 ri4/AtRZF1 OX) were used for further physiological characterizations.

To generate constitutively expressing lines, FL AtUAP1 cDNA was amplified by PCR using a gene-specific primer set (Supplementary Table S1). PCR products were initially cloned into pDONR/ZEO vector and confirmed by sequencing. Resultant plasmids were then directly cloned into a plant expression vector pGWB514 (Nakagawa et al. 2007) using a Gateway system (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s instruction. After that, the construct was transformed into WT and atrzf1 plants using A. tumefaciens strain GV3101 via in planta vacuum infiltration. T₂ transformants (hygromycin resistance) were selected from transgenic lines segregated as a single locus. T₃ homozygous AtUAP1-OX transgenic plants (OX3–4 and OX4–1) and AtUAP1-overexpression/atrzf1 lines (AtUAP1 OX2/atrzf1 and AtUAP1 OX17/atrzf1) were selected for phenotypic characterization.

To generate the AtRZF1-OX (AtRZF1 OX) construct, FL of AtRZF1 cDNA was amplified by PCR using a gene-specific primer set (Supplementary Table S1). PCR products were initially cloned into pDONR/ZEO vector and confirmed by sequencing. Resultant plasmids were then directly cloned into plant expression vector pGWB514 using the Gateway system (Invitrogen) according to the manufacturer’s instruction. This construct was transformed into WT plants using A. tumefaciens strain GV3101 via in planta vacuum infiltration. T₂ transformants (hygromycin resistance) were selected from transgenic lines segregated as a single locus. T₃ homozygous lines from 10 independent transgenic lines were obtained and one line with a high transgenic expression level was selected for phenotypic characterization.

Phenotype analysis and stress tests

To test response to osmotic stress and ABA, seeds were sown on half-strength MS medium supplemented with 400 mM mannitol and 1 μM ABA, respectively, and grown in a growth chamber. Cotyledon greening of each seedling was measured at 14 days. Experiments were done in three biological replicates with 50 or 60 seedlings per genotype per replicate.

Relative water content measurement

To obtain relative water content values, fourth or fifth rosette leaves of the same growth stage were excised and placed on open-lid Petri dishes at room temperature with a relative humidity of 60% under a dim light. Weights of rosette leaves were measured at various times. Leaf water content was described as percentage of initial FW.

Pro content determination

Pro content was measured as previously described (Bates et al. 1973). Briefly, Pro was extracted from 100 mg of plant leaves after grinding leaves in 1 ml of 3% sulfosalicylic acid. Then 200 µl of extract was reacted with 100 μl of ninhydrin reagent mixture (80% glacial acetic acid, 6.8% phosphoric acid and 70.17 mM ninhydrin) at 100°C for 60 min. The reaction was terminated by soaking in an ice bath. The reaction mixture was extracted with 200 μl of toluene and vortexed. Absorbance of the toluene layer was read at a wavelength of 520 nm using a ultraviolet/visible spectrophotometer (JASCO, Tokyo, Japan). Pro concentration was extrapolated from a standard curve and calculated on a FW basis as follows: [(ng Pro/ml × ml extraction buffer)/115.5 ng nmol]/g sample = nmol Pro/g FW material.

Statistical analysis

All statistical analyses were performed using SPSS 23.0 software (IBM Co, Armonk, NY, USA), including one-way analysis of variance and Tukey’s multiple range test. Different letters indicate statistically significant differences at P < 0.05 by Tukey’s multiple range test.

Supplementary Data

Supplementary data are available at PCP online.

Data Availability

Gene and protein sequences from this article can be found in the GenBank database (https://www.ncbi.nlm.nih.gov/) or The Arabidopsis Information Resource (TAIR) (https://www.arabidopsis.org/) under the following accession numbers: AtRZF1(At3g56580),AtAtUAP1(At5g14540),AtUAP2(At3g01560),UFD2(U22154),ZmUAP1(NP_001147786) and OsUAP1(XP_015648026).

Funding

Next-Generation BioGreen21 project (SSAC, PJ013171) and New Breeding Technology program (PJ01477701) funded by the Rural Development Administration, Republic of Korea, to [C.S.K].

Acknowledgements

J.H.M. and C.S.K. designed the study and interpreted study results. J.S.C. provided technical assistance with yeast functional complement analysis. J.H.M., C.R.P. and J.S.C. carried out interpretation and analysis of experimental results. J.H.M. and C.S.K. wrote the manuscript. All authors approved the final manuscript.

Disclosures

The authors have no conflict of interest to disclose.