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. 2019 Aug 8;20(11):1566–1573. doi: 10.1111/mpp.12855

The proteasome regulator PTRE1 contributes to the turnover of SNC1 immune receptor

Karen Thulasi Devendrakumar 1,†, Charles Copeland 1,2,†, Xin Li 1,✉
PMCID: PMC6804346  PMID: 31393057

Summary

Plants have evolved a sophisticated immune system in order to recognize and respond to microbes in their environments. Nucleotide‐binding leucine‐rich repeat (NLR) proteins detect the presence of specific effector molecules delivered into host cells by pathogens and activate strong defence responses. However, as excessive accumulation of NLRs can result in inappropriate immune responses, their abundance must be tightly regulated. Targeted degradation of NLRs through the ubiquitin proteasome pathway is an important mechanism to limit NLR accumulation. Mutations that perturb NLR degradation can cause autoimmune phenotypes. In this study, we show that the proteasome regulator PTRE1 also contributes to NLR degradation. ptre1 mutant plants exhibit increased defence marker gene expression and enhanced disease resistance against virulent pathogens. The stability of the NLR, SUPPRESSOR OF npr1‐1 CONSTITUTIVE 1 (SNC1) is also increased in the ptre1 mutant. Although the mouse homologue of PTRE1 was reported to interact with a Cell Division Control protein 48 (CDC48) homologue in vitro (Clemen et al., 2015), we only observed interaction between PTRE1 and AtCDC48A in a split luciferase assay, but not in co‐immunoprecipitation. In addition, a related Arabidopsis protein PTRE1h shares partial redundancy with PTRE1. Together, PTRE1 acts as a negative regulator of plant immunity partly by facilitating the degradation of immune receptors such as SNC1.

Keywords: NLR, plant immunity, proteasome, protein homeostasis, PTRE1, SNC1

Like all other multicellular organisms, plants face constant threats from pathogens such as bacteria and fungi in their environment. However, they have evolved a sophisticated innate immune system that enables them to recognize most potential pathogens and mount an immune response, preventing diseases (Jones and Dangl, 2006; Jones et al., 2016). Receptors on the plant cell surface can recognize elicitor molecules, which are often widely conserved among major groups of microbes (Jones and Dangl, 2006) and termed pathogen‐associated molecular patterns (PAMPs). PAMPs activate an immune response known as PAMP‐triggered immunity (PTI). However, many plant pathogens produce effector proteins which can disrupt PTI signalling, allowing an infection to be established. In turn, plants employ Resistance (R) proteins, which provide a second layer of pathogen detection by perceiving the intracellular effector proteins or their effects in the cell. Most R proteins belong to the nucleotide‐binding leucine‐rich repeat (NLR) family (Maekawa et al., 2011).

NLR activation leads to rapid and strong defence responses, including the production of reactive oxygen species, production of antimicrobial compounds and induction of a type of programmed cell death known as the hypersensitive response (HR). While these defence outputs are quite effective at preventing pathogen spread, they are detrimental to plant growth. Such negative effects are obvious in mutants with mutations that cause constitutive defence signalling, which results in severe dwarfism (Li et al., 2001; Wang et al., 2013; Yang et al., 2010). Thus, in the absence of pathogen attack, negative regulation of NLRs and their downstream signalling components is essential for optimal plant growth.

One important negative regulatory mechanism in eukaryotes is through targeted protein degradation by the ubiquitin proteasome system (UPS). In the UPS pathway, substrates are marked for degradation by the addition of the small protein ubiquitin (Vierstra, 2009). Ubiquitin can be conjugated onto lysine residues within the substrate itself, or onto a previously added ubiquitin moiety, forming a polyubiquitin chain. Polyubiquitinated substrates can subsequently be recognized by the 26S proteasome, a large protein complex with proteolytic function. The 26S proteasome is made up of the 19S regulatory particle, which recognizes the polyubiquitin conjugates and unfolds them, and the 20S core particle (CP), the proteolytic core that possesses multiple subunits with protease activity. The UPS system is critical in regulating almost all biological processes in higher plants, including immunity. Many regulatory proteins in plant defence, including PTI and NLR immune receptors, have been found to be under the control of the UPS system to guarantee an optimum immune output.

The Mutant, snc1 enhancing (MUSE) screen is a forward genetic screen for mutants that enhances the autoimmunity mediated by a gain‐of‐function NLR mutant suppressor of npr1-1 constitutive 1 (snc1), with the aim of identifying novel negative regulators of immunity (Huang et al., 2013). A number of the muse mutants identified in this screen are impaired in the turnover of the SNC1 protein by the UPS (Cheng et al., 2011; Copeland et al., 2016; Huang et al., 2014a, 2014b, 2016; Xu et al., 2015b). For example, a partial loss‐of‐function mutant of the Arabidopsis thaliana ATPase, Cell Division Control protein 48 homologue A (AtCDC48A) causes increased SNC1 protein accumulation, which is associated with enhanced disease resistance and dwarf, curly‐leaf morphology (Copeland et al., 2016). CDC48 homologues function in UPS‐mediated degradation of diverse substrates and are essential genes in eukaryotes, including Arabidopsis, yeast and animals (Baek et al., 2013; Dai and Li, 2001; Meyer et al., 2012; Rancour et al., 2002).

The best characterized function of CDC48 in the UPS is promoting the degradation of substrates by facilitating their recognition by the proteasome. A number of other proteins also assist in UPS‐mediated degradation by shuttling polyubiquitinated substrates to the proteasome, modulating proteasome activity (Yu and Matouschek, 2017). PI31 (also called PSMF1) has been reported to modulate proteasome activity in animal cells. While PI31 was shown to inhibit 20S CP activity in vitro, over‐expression of PI31 in the cell appears to increase the activity of the fully assembled 26S proteasome (Clemen et al., 2015; Li et al., 2014). In Arabidopsis, the PI31 homologue PTRE1 also positively regulates cellular proteasome activity (Yang et al., 2016). Upon perception of the plant hormone auxin, proteasome activity in wild‐type plants is reduced, while ptre1 mutants have constitutively lower proteasome activity that is unaffected by auxin. This suggests that a reduction in PTRE1 function is responsible for auxin‐mediated proteasome suppression. As a general regulator of 26S proteasome, it is not surprising that ptre1 plants also display a dwarf, curly‐leaf morphology similar to many mutants with constitutive immune signalling, including Atcdc48A‐4 (Fig. 1A) (Copeland et al., 2016; Yang et al., 2016). This striking resemblance between ptre1 and Atcdc48A‐4 plants prompted us to examine whether PTRE1 also plays a role in immune regulation.

Figure 1.

Figure 1

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PTRE1 negatively regulates immunity. (A) Morphology of soil‐grown 5‐week‐old Arabidopsis thaliana plants of the indicated genotypes. (B) Growth of the bacterial pathogen Pseudomonas syringae pv. maculicola (P.s.m.) ES4326 on plants of the indicated genotypes. Plants were infiltrated with bacterial suspension (OD600 = 0.001) and bacterial titre was measured after 0 and 3 days. Error bars represent standard deviation (n = 3). The letters indicate significant difference between the different genotypes as determined using a Tukey HSD test. Genotypes denoted using different letters have significant difference (P < 0.01). The experiment was performed independently three other times with similar results. (C) Growth of Hyaloperonospora arabidopsidis (H.a.) Noco2 on plants of the indicated genotypes. Plants were approximately 2.5 weeks old when spray‐inoculated with spore solution of the oomycete (105 spores/mL). Spores were counted 1 week later. Error bars represent standard error of the mean (SEM). The asterisk indicates a significant difference (P < 0.05) from Col, as determined by a t‐test. (D–F) PR1 (D), PR2 (E) and ICS1 (F) gene expression in the indicated genotypes. qRT‐PCR was performed using RNA extracted from 4‐week‐old plants grown on soil. The letters indicate significant differences between the different samples as determined using a Tukey HSD test. Samples denoted using different letters have a significant difference (P < 0.01). The error bars represent standard deviation of the technical replicates (n = 3). The experiment was performed two other times with similar results.

In order to determine whether the ptre1 morphological phenotype is associated with increased immune signalling, we performed infection assays using the virulent bacterial pathogen Pseudomonas syringae pv. maculicola (P.s.m.) ES4326 and an oomycete pathogen Hyaloperonospora arabidopsidis (H.a.) Noco2 according to the protocols described by Li et al. (2001). Enhanced disease resistance was observed when ptre1 mutant plants were challenged with both pathogens (Fig. 1B,C). We also examined expression of the defence marker genes PATHOGENESIS RELATED 1 (PR1), PR2 and ICS1 (Isochorismate synthase 1) using qRT‐PCR. ptre1 plants indeed also showed increased expression of these genes (Fig. 1D–F). Taken together, knocking out PTRE1 leads to autoimmunity similar to that in Atcdc48A‐4. However, the ptre1 mutant is not as temperature sensitive as the other autoimmune mutants, such as snc1 or Atcdc48A‐4 (Fig. S1). This is likely because PTRE1 is a general proteasome regulator and knocking out PTRE1 is expected to yield other growth and developmental defects unrelated to immunity.

Atcdc48A‐4 and a number of other muse mutants have reduced turnover of NLR proteins such as SNC1, causing constitutive immune signalling. Since PTRE1 is thought to promote protein turnover by the proteasome, and may interact with AtCDC48A (Clemen et al., 2015), we first tested whether ptre1 mutants exhibit altered SNC1 levels. Total protein was extracted according to Huang et al. (2014b), and immunoblotted using a SNC1‐specific antibody (Li et al., 2010). SNC1 protein abundance was significantly higher in ptre1 plants than in the wild‐type (WT) (Fig. 2A,B). We did not observe increased SNC1 gene expression in the ptre1 plants, suggesting that the increased SNC1 protein abundance is due to increased protein stability (Fig. 2C). To further confirm that the increase in SNC1 protein level is not due to indirect effects of increased immunity in the ptre1 mutant, we introduced the ptre1 mutation into the eds1 background, which is impaired in feedback up‐regulation of NLR gene transcription (Wiermer et al., 2005). The ptre1 eds1 double mutant was morphologically similar to the ptre1 single mutant (Fig. 2D), likely due to pleiotropic effects such as the auxin‐related phenotypes reported by Yang et al. (2016). A significant increase in SNC1 protein abundance was still observed in the ptre1 eds1 double mutant plants, further supporting our hypothesis that PTRE1 promotes SNC1 turnover (Fig. 2E,F).

Figure 2.

Figure 2

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Knockout mutant of PTRE1 displays increased SNC1 protein stability. (A) SNC1 protein level in the indicated genotypes. Total protein was extracted from 3‐week‐old Arabidopsis thaliana plants grown on soil and immunoblot analysis was performed using a SNC1‐specific antibody (Li et al., 2010). Ponceau staining is shown as a loading control. (B) Quantification of SNC1 protein level from repeats of experiments as described in (A). For each sample, the intensity of the SNC1 band was normalized to the Ponceau staining, then the relative intensity in Col was set to 1. Error bars represent standard error of the mean. Asterisks indicate a significant difference (P < 0.05) from Col as determined by a t‐test. The experiment was repeated at least three times with similar results. (C) SNC1 gene expression in the indicated genotypes. qPCR was performed on RNA extracted from plants in (A). SNC1 expression was normalized to ACTIN7. Error bars represent standard error of the mean. (D) Morphology of 4‐week‐old soil‐grown plants of the indicated genotypes. (E) SNC1 protein level in the indicated genotypes. Experiment was performed as in (A). (F) Quantification of SNC1 protein level from the repeats of experiments as described in (E). (G) Morphology of snc1 mutants over‐expressing PTRE1. PTRE1 driven by the 35S promoter was introduced into the snc1 background. The image shows plants from six independent single copy transgenic lines. (H) PTRE1 expression levels in the PTRE1 over‐expression (OE) lines in the snc1 background (as in G). Homozygous T3 plants were used for the PTRE1 expression level quantification. qRT‐PCR was performed using RNA extracted from 4‐week‐old plants grown on soil. The PTRE1 expression level is set to 1 for snc1. The error bars represent standard deviation of the technical replicates (n = 3).

As PTRE1 negatively regulates the level of SNC1 protein, we wondered if the levels of other NLRs are also affected. Therefore, we introduced the ptre1 mutation, by crossing, into lines expressing either RPM1‐myc (Boyes et al., 1998) or RPS4‐HA (Wirthmueller et al., 2007). Neither NLR showed increased abundance in the ptre1 background compared to the parental line (Fig. S2A). While it is possible that loss of PTRE1 causes a change in RPM1 or RPS4 levels that is undetectable in an immunoblot, this result suggests that PTRE1 displays some levels of specificity in regulating protein turnover, even within the NLR family.

The exact function of PTRE1 in proteasome regulation is still unclear (Fort et al., 2015). Some genetic and biochemical evidence indicates that PTRE1 homologues stimulate 26S proteasome activity. Over‐expression of Drosophila PI31 suppresses phenotypes caused by mutant proteasome subunits with impaired function (Bader et al., 2011). Yang et al. (2016) reported that over‐expression of PTRE1 in Arabidopsis resulted in reduced accumulation of IAA proteins. Therefore, we tested whether PTRE1 over‐expression could decrease SNC1 stability. A 35S::PTRE1 over‐expression construct was made and 35S::PTRE1 could complement the ptre1 mutant morphological phenotype and bring down the SNC1 protein level to near wild‐type level (Fig. S2B,C), suggesting that 35S::PTRE1 is functional. Since wild‐type plants show a low steady‐state SNC1 level, over‐expression of PTRE1 would be unlikely to cause an observable protein level reduction. Instead, we introduced a 35S::PTRE1 over‐expression construct into the snc1 mutant, in which the dwarf morphology and increased immunity are due to increased SNC1 protein abundance (Cheng et al., 2011). If PTRE1 over‐expression reduces SNC1 stability, we would expect to see full or subtle suppression of the autoimmune phenotype in 35S::PTRE1 snc1 plants. The PTRE1 expression level in the over‐expression lines was quantified by qRT‐PCR and PTRE1 was found to be significantly over‐expressed in these transgenic lines (Fig. 2F). However, none of these lines showed any obvious morphological differences from the snc1 single mutant, suggesting that SNC1 turnover is not affected upon PTRE1 over‐expression (Fig. 2G). It is possible that PTRE1 is not the rate‐limiting component in the degradation of SNC1. Similarly, no general effect was seen on the degradation of known UPS targets when PI31 was over‐expressed in mammalian cells (Li et al., 2014; Zaiss et al., 2002).

The ptre1 eds1 was further studied and its immune phenotype was characterized. In an infection assay using P.s.m. ES4326, the bacterial growth in the ptre1 eds1 double mutant was comparable to the eds1 single mutant (Fig. 3A). This further shows that the increased resistance of ptre1 is due to immunity‐related changes rather than indirect non‐specific pleiotropic effects of the ptre1 mutation. Consistently, PR1, PR2 and ICS1 gene expression levels are lower in the ptre1 eds1 double mutant compared to the ptre1 single mutant (Fig. 3B–D).

Figure 3.

Figure 3

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eds1 suppresses the enhanced immunity of the ptre1 mutant. (A) Growth of the bacterial pathogen Pseudomonas syringae pv. maculicola (P.s.m.) ES4326 in leaves of Arabidopsis thaliana plants of the indicated genotypes. Plants were infiltrated with bacterial suspension (OD600 = 0.001) and bacterial titre was measured after 0 and 3 days. Error bars represent sample standard deviation (n > 2). The letters indicate significant difference between the different samples as determined using a Tukey HSD test. Samples denoted using different letters have significant difference (P < 0.05). The experiment was performed independently three other times with similar results. (B–D) PR1 (B), PR2 (C) and ICS1 (D) gene expression in the indicated genotypes. qRT‐PCR was performed using RNA extracted from 4‐week‐old plants grown on soil. The letters indicate significant difference between the different samples as determined using a Tukey HSD test. Samples denoted using different letters have significant difference (P < 0.01). The error bars represent standard deviation of the technical replicates (n = 3). The experiment was performed two other times with similar results.

The molecular mechanism of PTRE1 function remains to be elucidated. PI31 interacts with F‐box proteins in both Drosophila and humans (Bader et al., 2011; Kirk et al., 2008). However, at least in Drosophila this interaction seems to stabilize PI31 but is not strictly required for its proteasome‐regulatory function. In addition, the murine homologues of AtCDC48A and PTRE1 (VCP and PMSF1, respectively) show a direct interaction in vitro, which modulates their ability to regulate the activity of purified 20S proteasome core particles (Clemen et al., 2015). Since Arabidopsis PTRE1 and AtCDC48A are both thought to positively regulate 26S proteasome activity, we hypothesized that they may also interact in planta. Therefore, we tested for protein–protein interactions using a co‐immunoprecipitation (co‐IP) assay, which was carried out as described by Xu et al. (2015a). PTRE1 fused to a C‐terminal flag epitope tag (PTRE1‐flag) and AtCDC48A with a C terminal GFP tag (AtCDC48A‐GFP) (Copeland et al., 2016) were transiently co‐expressed in Nicotiana benthamiana using Agrobacterium‐mediated transformation at an OD600 of 0.3 for each strain, with a strain containing an empty vector being used as negative control. Infiltrated leaf tissue was harvested after 48 h, and α‐flag conjugated to agarose beads was used for the co‐IP. As shown in Fig. 4A, AtCDC48A‐GFP did not co‐IP with PTRE1‐flag. However, it is still possible that PTRE1 may work together with AtCDC48A through a weak or transient association that cannot be detected by co‐IP.

Figure 4.

Figure 4

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PTRE1 does not co‐immunoprecipitate with AtCDC48A but interacts with it in the split luciferase assay. (A) AtCDC48A‐GFP was co‐expressed in Nicotiana benthamiana leaves with either PTRE1‐flag or with an empty vector, and the total protein was extracted after 48 h. The protein was subjected to immunoblot analysis using the indicated antibodies. Arrowheads show the size of the prospective bands and the approximate protein sizes are indicated to the right of the blots. Ponceau staining of the blot is shown as a loading control. (B) Cartoon showing the infiltration areas on the N. benthamiana leaf for the split luciferase assay. The legend denotes the gene constructs in the Agrobacterium infiltrated in that area. (C) Luminescence recorded after infiltration of 1 mM luciferin into the Agrobacterium infiltration sites. The heat map scale is on the bottom with dark red representing high levels of luminescence and blue representing low levels of luminescence. (D) False colour image of the same leaf photographed under white light showing the infiltration areas.

The interaction between the two proteins was further examined using the split luciferase complementation assay. PTRE1 and AtCDC48A were cloned into a modified pCambia1300 vector with the CaMV 35S promoter. PTRE1 was fused to the N‐terminal part of the luciferase protein at its C terminus (PTRE1‐NLuc). AtCDC48A was fused to the C‐terminal fragment of luciferase at its C terminus (AtCDC48‐CLuc). These constructs were then transformed into Agrobacterium strain GV3101 for infiltration into the leaves of N. benthamiana. The Agrobacterium strains carrying different vectors were infiltrated using a syringe into the leaves in combinations as seen in Fig. 4B with each of the Agrobacterium strains diluted to a final OD600 of 0.2 with the same infiltration solution as used in the co‐IP experiment. The MPK4‐NLuc and CLuc‐MKK6 interaction serves as a positive control. Luminescence was observed in the areas of the leaf infiltrated with the positive control and the area infiltrated with PTRE1‐NLuc and AtCDC48‐CLuc (Fig. 4C,D), demonstrating a direct protein–protein interaction between the two proteins. The reason we did not see interaction in co‐IP but observed interaction in the split luciferase assay could be because the two proteins have a weak and/or transient interaction that is difficult to observe with co‐IP. Overall, a more thorough examination of the genetic and biochemical interaction network of PTRE1 is required to understand the mechanism by which it regulates proteasome activity.

A BLAST search of the Arabidopsis genome using PTRE1 as a query revealed a homologous sequence, AT1G48530, which we named PTRE1h. Based on an alignment of PTRE1 and PTRE1h, PTRE1h is a truncated sequence, containing homology to the first three exons of PTRE1 (Fig. S3A). However, the deduced amino acid sequences of both genes within this region share 56% identity and 70% similarity, prompting us to test whether PTRE1h may share redundant function with PTRE1. A ptre1h mutant containing an exonic T‐DNA insertion was morphologically indistinguishable from Col (Fig. S3B). To test for possible redundancy between PTRE1 and PTRE1h, a ptre1 ptre1h double mutant was generated. The morphological phenotype of the double mutant was not altered compared to the ptre1 single mutant, nor were double mutants more resistant to H.a. Noco2 or P.s.m ES4326 infection (Fig. S3C,D). However, defence markers PR1, PR2 and the ICS1 expression levels are consistently higher in the ptre1 ptre1h double mutant as compared to those in the ptre1 single mutant (Fig. S3E–G), suggesting that PTRE1h is partially redundant with PTRE1. The difference in immunity between ptre1 and ptre1 ptre1h could be too small to be detected using the infection experiments that we performed. When the SNC1 protein level was quantified in the ptre1 and ptre1 ptre1h mutants to see whether the ptre1h mutation further contributed to SNC1 protein stability, we could observe an overall increase in the SNC1 protein level in the ptre1 ptre1h double mutant compared to the single mutant (Fig. S4A). However, this difference was not statistically significant among repeats of the experiment (Fig. S4B).

NLR turnover through the UPS is crucial to limit NLR over‐accumulation and prevent autoimmunity (Holt et al., 2005; Li et al., 2015). In this study, we show that the proteasome regulator PTRE1 is a negative regulator of immunity and promotes SNC1 turnover. The ptre1 mutant displays a dwarf phenotype, increased defence marker gene expression and enhanced resistance to virulent pathogens (Fig. 1). SNC1 protein abundance is also increased in the ptre1 mutant. The Arabidopsis genome contains a truncated homologue of PTRE1, PTRE1h, that shares high homology in the N‐terminal region (Fig. S3). PTRE1h seems to be partially redundant with PTRE1.

The ptre1 mutant displays a range of defects potentially related to auxin, abscisic acid and brassinosteroid signalling, suggesting that PTRE1 promotes the turnover of many different substrates (Yang et al., 2016). Our finding in this study that PTRE1 is involved in SNC1 turnover adds to the previously identified model of NLR degradation by identifying another component that likely acts together with the proteasome and AtCDC48A. However, we still do not know the molecular events involved in PTRE1 regulation of proteasome activity. A more thorough characterization of protein interactors of PTRE1 will be required to determine how it facilitates the degradation of specific substrates.

Supporting information

Fig. S1 ptre1 morphological phenotype is not suppressed when grown at high temperature. The figure shows the plants of the indicated genotypes grown at the indicated temperatures.

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Fig. S2 PTRE1 does not affect RPM1 and RPS4 levels, and PTRE1 over‐expression is able to complement the phenotype of the ptre1 mutant. (A) Protein levels of RPM1‐myc (left) and RPS4‐HA (right) in the indicated genotypes. Total protein was extracted from 4‐week‐old plants grown on soil, and immunoblot analysis was performed using the respective antibodies. Ponceau staining of the blot is shown as a loading control. (B) Two T1 ptre1 plants (#1 and #2) over‐expressing PTRE1 are shown above with ptre1 and Col as controls. (C) SNC1 protein level in the complementation lines shown above. The Ponceau‐stained band is shown as the loading control.

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Fig. S3 PTRE1h is partially redundant with PTRE1. (A) Gene model of PTRE1 and PTRE1h. Exons are represented as boxes and introns are represented as lines. Arrowheads indicate the positions of the T‐DNA insertions in the mutants described. (B) Morphology of 5‐week‐old soil‐grown plants of the indicated genotypes. (C) Growth of Hyaloperonospora arabidopsidis. Noco2 on plants of the indicated genotypes. Three‐week‐old plants were inoculated with spore suspension (105  spores/mL) and grown at 18 °C for a week. Spores were counted 1 week later. Error bars represent sample standard deviation (n = 3). The letters indicate significant difference between the different samples as determined using a Tukey Honest Significant Difference (HSD) test. Samples denoted using different letters have significant difference (P < 0.05). The experiment was performed independently two other times with similar results. (D) Growth of the bacterial pathogen Pseudomonas syringae pv. maculicola ES4326 in leaves of plants of the indicated genotypes. Plants were infiltrated with bacterial suspension (OD600 = 0.001) and bacterial titre was measured after 0 and 3 days. Error bars represent sample standard deviation (n = 3). The letters indicate significant difference between the different samples as determined using a Tukey HSD test. Samples denoted using different letters have significant difference (P < 0.01). The experiment was performed independently three other times with similar results. (E–G) PR1 (E), PR2 (F) and ICS1 (G) gene expression in the indicated genotypes. qRT‐PCR was performed using RNA extracted from 4‐week‐old plants grown on soil. The letters indicate significant difference between the different samples as determined using a Tukey HSD test. Samples denoted using different letters have significant difference (P < 0.01). The experiment was performed two other times with similar results.

Click here for additional data file. (153.7KB, pdf)

Fig. S4 SNC1 protein levels in the ptre1 and the ptre1 ptre1h are not significantly different. (A) SNC1 protein levels quantified in 4‐week‐old soil‐grown Arabidopsis thaliana plants of the indicated genotypes using a SNC1‐specific antibody (Li et al., 2010). The Ponceau‐stained band is shown as a loading control. The experiment was repeated three times with similar results. (B) The SNC1 protein band intensities were quantified using ImageJ, normalized to the loading control band and then the relative intensity in Col was set to 1. The bar graph shows the mean normalized SNC1 band intensity of the three repeats of the experiment and the error bars denote the standard deviation of the normalized band intensity between experiments. The letters indicate the significant difference between the different samples as determined using a Tukey HSD test. Samples denoted using different letters have significant difference (P < 0.01).

Click here for additional data file. (166.7KB, pdf)

Method Plant growth condition and pathogen infection assays; quantitative RT‐PCR; protein extraction and immunoprecipitation.

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Acknowledgements

We thank Dr Jane Parker and Dr Jeff Dangl for kindly providing us with the RPS4‐HA and RPM1‐Myc transgenic lines. We also thank the Arabidopsis Biological Resource Center (ABRC) for providing seeds of the T‐DNA mutants. The research was supported by a Natural Science and Engineering Research Council of Canada (NSERC) Discovery grant to X.L. and an NSERC Vanier Canada Graduate Scholarship to C.C.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Fig. S1 ptre1 morphological phenotype is not suppressed when grown at high temperature. The figure shows the plants of the indicated genotypes grown at the indicated temperatures.

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Fig. S2 PTRE1 does not affect RPM1 and RPS4 levels, and PTRE1 over‐expression is able to complement the phenotype of the ptre1 mutant. (A) Protein levels of RPM1‐myc (left) and RPS4‐HA (right) in the indicated genotypes. Total protein was extracted from 4‐week‐old plants grown on soil, and immunoblot analysis was performed using the respective antibodies. Ponceau staining of the blot is shown as a loading control. (B) Two T1 ptre1 plants (#1 and #2) over‐expressing PTRE1 are shown above with ptre1 and Col as controls. (C) SNC1 protein level in the complementation lines shown above. The Ponceau‐stained band is shown as the loading control.

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Fig. S3 PTRE1h is partially redundant with PTRE1. (A) Gene model of PTRE1 and PTRE1h. Exons are represented as boxes and introns are represented as lines. Arrowheads indicate the positions of the T‐DNA insertions in the mutants described. (B) Morphology of 5‐week‐old soil‐grown plants of the indicated genotypes. (C) Growth of Hyaloperonospora arabidopsidis. Noco2 on plants of the indicated genotypes. Three‐week‐old plants were inoculated with spore suspension (105  spores/mL) and grown at 18 °C for a week. Spores were counted 1 week later. Error bars represent sample standard deviation (n = 3). The letters indicate significant difference between the different samples as determined using a Tukey Honest Significant Difference (HSD) test. Samples denoted using different letters have significant difference (P < 0.05). The experiment was performed independently two other times with similar results. (D) Growth of the bacterial pathogen Pseudomonas syringae pv. maculicola ES4326 in leaves of plants of the indicated genotypes. Plants were infiltrated with bacterial suspension (OD600 = 0.001) and bacterial titre was measured after 0 and 3 days. Error bars represent sample standard deviation (n = 3). The letters indicate significant difference between the different samples as determined using a Tukey HSD test. Samples denoted using different letters have significant difference (P < 0.01). The experiment was performed independently three other times with similar results. (E–G) PR1 (E), PR2 (F) and ICS1 (G) gene expression in the indicated genotypes. qRT‐PCR was performed using RNA extracted from 4‐week‐old plants grown on soil. The letters indicate significant difference between the different samples as determined using a Tukey HSD test. Samples denoted using different letters have significant difference (P < 0.01). The experiment was performed two other times with similar results.

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Fig. S4 SNC1 protein levels in the ptre1 and the ptre1 ptre1h are not significantly different. (A) SNC1 protein levels quantified in 4‐week‐old soil‐grown Arabidopsis thaliana plants of the indicated genotypes using a SNC1‐specific antibody (Li et al., 2010). The Ponceau‐stained band is shown as a loading control. The experiment was repeated three times with similar results. (B) The SNC1 protein band intensities were quantified using ImageJ, normalized to the loading control band and then the relative intensity in Col was set to 1. The bar graph shows the mean normalized SNC1 band intensity of the three repeats of the experiment and the error bars denote the standard deviation of the normalized band intensity between experiments. The letters indicate the significant difference between the different samples as determined using a Tukey HSD test. Samples denoted using different letters have significant difference (P < 0.01).

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Method Plant growth condition and pathogen infection assays; quantitative RT‐PCR; protein extraction and immunoprecipitation.

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