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. 2016 Dec 6;28(12):3038–3051. doi: 10.1105/tpc.16.00574

Orosomucoid Proteins Interact with the Small Subunit of Serine Palmitoyltransferase and Contribute to Sphingolipid Homeostasis and Stress Responses in Arabidopsis[OPEN]

Jian Li 1,1, Jian Yin 1,1, Chan Rong 1,1, Kai-En Li 1,1, Jian-Xin Wu 1,1, Li-Qun Huang 1, Hong-Yun Zeng 1, Sunil Kumar Sahu 1, Nan Yao 1,2
PMCID: PMC5240739  PMID: 27923879

Loss of function of the orosomucoid proteins stimulates the de novo biosynthesis of sphingolipids, causes an early senescence phenotype, and enhances resistance to oxidative stress and pathogen infection.

Abstract

Serine palmitoyltransferase (SPT), a pyridoxyl-5′-phosphate-dependent enzyme, catalyzes the first and rate-limiting step in sphingolipid biosynthesis. In humans and yeast, orosomucoid proteins (ORMs) negatively regulate SPT and thus play an important role in maintaining sphingolipid levels. Despite the importance of sphingoid intermediates as bioactive molecules, the regulation of sphingolipid biosynthesis through SPT is not well understood in plants. Here, we identified and characterized the Arabidopsis thaliana ORMs, ORM1 and ORM2. Loss of function of both ORM1 and ORM2 (orm1 amiR-ORM2) stimulated de novo sphingolipid biosynthesis, leading to strong sphingolipid accumulation, especially of long-chain bases and ceramides. Yeast two-hybrid, bimolecular fluorescence complementation, and coimmunoprecipitation assays confirmed that ORM1 and ORM2 physically interact with the small subunit of SPT (ssSPT), indicating that ORMs inhibit ssSPT function. We found that orm1 amiR-ORM2 plants exhibited an early-senescence phenotype accompanied by H2O2 production at the cell wall and in mitochondria, active vesicular trafficking, and formation of cell wall appositions. Strikingly, the orm1 amiR-ORM2 plants showed increased expression of genes related to endoplasmic reticulum stress and defenses and also had enhanced resistance to oxidative stress and pathogen infection. Taken together, our findings indicate that ORMs interact with SPT to regulate sphingolipid homeostasis and play a pivotal role in environmental stress tolerance in plants.

INTRODUCTION

In eukaryotes, sphingolipids make up ∼40% of the lipids of the plasma membrane and are also abundant in other endomembranes. The functions of these key lipids have been intensively investigated in mammals and yeast for decades (Hannun and Obeid, 2008), and recent work has begun to explore sphingolipid biochemistry in plants. Sphingolipids play pivotal roles as membrane structural components, as bioactive molecules involved in signal transduction and cell regulation, and in a wide range of other biological processes, including secretion, programmed cell death, autophagy, stress responses, and cell-cell interactions (Liang et al., 2003; Markham et al., 2011; Sentelle et al., 2012; Bi et al., 2014; Li et al., 2015; Wu et al., 2015). On the outer leaflet of the membrane, sphingolipids form membrane microdomains with cholesterol to provide conformational support for membrane proteins and serve as a platform for recruitment of signaling molecules (Lingwood and Simons, 2010).

In humans, changes in sphingolipid contents have been closely linked to diabetes (Summers and Nelson, 2005), cancer (Modrak et al., 2006), Alzheimer’s disease (Mizuno et al., 2016), cardiovascular disease, and respiratory disease (Park et al., 2006). For example, sphingosine-1-phosphate can bind to the G protein-coupled receptor EDG5 to inhibit the activity of Rac protein, which could prevent the metastasis of tumor cells (Okamoto et al., 2000). The sphingosine-1-phosphate analog fingolimod (FTY720) has a similar activity and can be used for cancer treatment (Brinkmann et al., 2010). Sphingolipids also function in plant development and responses to biotic or abiotic stresses (zhen et al., 2006, 2009; Dietrich et al., 2008; Teng et al., 2008; Markham et al., 2011; Ternes et al., 2011; König et al., 2012; Zhang et al., 2013; Bi et al., 2014).

Regulation of the levels of sphingolipids involves the modulation of key enzymes such as ceramide synthases, ceramidases, ceramide kinase, glucosylceramidase, and inositolphosphorylceramidase (Liang et al., 2003; Wang et al., 2008; Ternes et al., 2011; Bi et al., 2014; Li et al., 2015; Msanne et al., 2015; Wu et al., 2015). Serine palmitoyltransferase (SPT) is a pyridoxyl-5′-phosphate-dependent enzyme that catalyzes the first and rate-limiting step in sphingolipid biosynthesis, the condensation between l-serine and a long-chain acyl thioester such as palmitoyl-CoA (C16-CoA) to generate long-chain bases (LCBs) (Chen et al., 2006). In Arabidopsis thaliana, the SPT enzyme has three subunits, LCB1, LCB2a, and LCB2b. The fbr11-2/lcb1-1 mutant, a loss-of-function mutation of LCB1, shows abnormal development, initiating apoptotic cell death in binucleate microspores, and the LCB2 loss-function mutant displays gametophytic lethality (Chen et al., 2006; Dietrich et al., 2008; Teng et al., 2008).

Work in animals and yeast showed that orosomucoid (ORM) proteins can modulate SPT activity (Breslow et al., 2010; Han et al., 2010; Gururaj et al., 2013). The ORM family proteins are endoplasmic reticulum (ER)-resident membrane proteins encoded by ORM or ORMDL genes, which are conserved from yeast to humans (Hjelmqvist et al., 2002; Moffatt et al., 2007). Depletion of the mammalian ORMDL1-3 eliminates the feedback of exogenous ceramide on ceramide biosynthesis, indicating that ORMDL proteins function as the primary regulators of ceramide biosynthesis in mammalian cells (Siow and Wattenberg, 2012). In yeast, genetic studies established a link between ORM1 and ORM2 and sphingolipid metabolism, as deletion of ORM1 and ORM2 leads to toxic accumulation of sphingolipids, whereas overexpression of ORM1 or ORM2 leads to reduced sphingolipid levels (Breslow et al., 2010). Thus, ORM1 and ORM2 negatively regulate sphingolipid synthesis.

Yeast (Saccharomyces cerevisiae) Orm proteins regulate sphingolipid metabolism by forming a multiprotein complex with SPT, termed the SPOTS complex, which also contains the SPT accessory subunit Tsc3 and the phosphoinositide phosphatase Sac1. Phosphorylation of Orm proteins, mediated by both branches of the TOR signaling pathway, regulates SPT activity to maintain sphingolipid homeostasis (Breslow et al., 2010; Han et al., 2010; Breslow, 2013). The Ypk1 protein kinase, downstream of rapamycin complex TORC, regulates Orm1 and Orm2 phosphorylation (Roelants et al., 2011; Berchtold et al., 2012; Niles et al., 2012; Sun et al., 2012; Shimobayashi et al., 2013). Orm proteins’ SPT-inhibitory activity is subject to feedback regulation by multiple sphingolipid intermediates, including LCBs, ceramide, and complex sphingolipids. When sphingolipid biosynthesis is disrupted, phosphorylation of their amino termini activates Orm1 and Orm2, thus enabling a compensatory increase in SPT activity (Breslow et al., 2010; Liu et al., 2012). Orm1 activity is adjusted in response to manipulation of Orm2 expression levels, with increased Orm2 expression causing a corresponding increase in Orm1 phosphorylation and vice versa. Phosphoregulation of Orm proteins controls sphingolipid biosynthesis in response to various stresses, including heat stress, ER stress, iron stress, and cell wall stress (Sun et al., 2012; Lee et al., 2012; Gururaj et al., 2013).

In addition to ORM, eukaryotes have a class of polypeptides, the SPT small subunits (ssSPT), which can regulate SPT activity. For example, yeast has an 80-amino acid polypeptide called Tsc3, which combines with the LCB1 and LCB2 subunits to form a trimer, thereby activating SPT (Gable et al., 2000). Humans have two ssSPT: ssSPTa (68 amino acids) and ssSPTb (76 amino acids). Coexpression of human ssSPTa or ssSPTb and LCB1, LCB2a, or LCB2b can activate SPT in yeast lcb1 lcb2△ mutants (Han et al., 2009). Recent work reported that Arabidopsis has two ssSPTs, encoded by ssSPTa (At1g06515) and ssSPTb (At2g30942). These 56-amino acid ssSPTs can activate SPT and play an important role in the formation of mature pollen (Kimberlin et al., 2013).

Considerable progress has been made in advancing our knowledge of plant sphingolipid metabolism in the past 10 to 15 years. However, our understanding of ORMs, particularly in plants, remains in its infancy. Here, we reported functional characterization of the ORM genes (ORM1, AT1G01230; ORM2, AT5G42000) in Arabidopsis. We discovered that ORM can interact with and inhibit ssSPT, thus affecting sphingolipid levels. Our studies strongly suggest that ORM plays a key role in the plant response to biotic and abiotic stress.

RESULTS

Loss of ORM1 and ORM2 Function Causes an Early-Senescence Phenotype and High Sphingolipid Accumulation

ORM family genes encode conserved ER membrane proteins in eukaryotic cells. Arabidopsis has two genes encoding homologs of S. cerevisiae Orm, AT1G01230 and AT5G42000, designated ORM1 and ORM2 (Figure 1A), which encode 157- and 154-amino acid polypeptides with 39 and 35% identity to Orm1 in yeast, respectively (Supplemental Figure 1A). Using the TMHMM protein structure analysis tool (http://www.cbs.dtu.dk/services/TMHMM/), we found that ORM1 and ORM2 have three predicted transmembrane domains (Supplemental Figure 2). The N-terminal extension found in yeast was absent from Arabidopsis ORMs, but phosphorylation site analysis (using NetPhos and the plant-specific algorithm PhosPhAt) showed ORM1 and ORM2 to have possible phosphorylation sites (Supplemental Figure 1A). ORM1 and ORM2 were expressed at higher levels in siliques than in other tissues (Supplemental Figure 1B). Our confocal microscopy observations confirmed that ORM1 and ORM2 localized to the ER of Arabidopsis (Figure 1D).

Figure 1.

Figure 1.

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Loss of Function of ORM1 and ORM2 Causes an Early-Senescence Phenotype and Sphingolipid Accumulation.

(A) The structures of Arabidopsis ORM1 and ORM2 and the locations of T-DNA insertions.

(B) and (C) The phenotype of wild-type and ORM loss-of-function plants at 26 (B) and 35 (C) d old. The white arrows indicate leaf senescence in orm1 amiR-ORM2 plants.

(D) Subcellular localization of ORM proteins. The fusion constructs 35Spro:GFP-ORM1 and 35Spro:GFP-ORM2 were coexpressed with the ER mCherry marker (TAIR, CD3-960) by transient expression in protoplasts. The 35Spro:GFP construct and CD3-960 were cotransformed as controls. The images were obtained by confocal microscopy after 16 h incubation. The experiments were repeated at least three times with similar results. Bars = 5 μm.

(E) Sphingolipids accumulated in orm1 amiR-ORM2 plants. Sphingolipids were extracted from 28-d-old plants following the steps described in Methods. The main sphingolipids were separated and identified by HPLC-ESI-MS. The amount of total LCBs, ceramides (Cer), hydroxyceramides (hCer), and glucosylceramides (gCer) was quantified (see Supplemental Figure 4 for major LCB and ceramide species). The experiment was repeated three times with similar results using independent samples. Values are means ± se from three technical replicates. Asterisks show a significant difference from the wild type using Student’s t test (***P < 0.001).

To test the function of the Arabidopsis ORMs, we characterized the available T-DNA mutant lines for these genes: SALK_046054 (predicted T-DNA insertion in the first intron of ORM1) and SAIL_1286_D09 (predicted T-DNA insertion in the 5′ untranslated region of ORM2). Homozygous lines were identified for each of these mutants. We did not detect the full-length transcript of ORM1 in SALK_046054, indicating that the line is a null mutant (Supplemental Figures 1C and 1E). In Arabidopsis, according to TAIR (http://www.arabidopsis.org), ORM2 has two splice isoforms, AT5G42000.1 and AT5G42000.2. However, our RT-PCR experiments detected only the transcripts of AT5G42000.1 (ORM2.1) in plants (Supplemental Figure 1D). Since the full-length transcript of ORM2 was still detected in SAIL_1286_D09 (Supplemental Figure 1D), we also generated an artificial microRNA (amiR) line targeting ORM2 (amiR-ORM2) (Supplemental Figure 1E). For subsequent analysis, we crossed orm1 with amiR-ORM2 plants and obtained the orm1 amiR-ORM2 homozygous lines. In addition, we placed ORM1 or ORM2 cDNAs under control of the CaMV35S promoter to create the overexpression lines ORM1-OX and ORM2-OX in Arabidopsis (Supplemental Figures 1C and 1D). The orm1 amiR-ORM2 plants initially underwent the same development as the wild type (Supplemental Figure 3A, top panel; orm1 amiR-ORM2-6 and orm1 amiR-ORM2-5). However, in late development, the orm1 amiR-ORM2 plants showed accelerated senescence, characterized by early chlorosis of rosette leaf tips (Figures 1B and 1C). By contrast, the development of the orm1 single mutant and amiR-ORM2 plants was similar to that of wild-type plants (Supplemental Figure 3A).

In yeast, Orm proteins act as negative mediators of sphingolipid biosynthesis by inhibiting SPT activity (Han et al., 2010). To determine the role of ORM1 and ORM2 in the regulation of sphingolipid biosynthesis in Arabidopsis, we comprehensively analyzed the major classes of sphingolipids in orm1 and amiR-ORM2 plants. We observed no obvious changes in sphingolipids in orm1 and amiR-ORM2 plants compared with the wild type (Supplemental Figure 3B). However, the sphingolipid profile of 28-d-old orm1 amiR-ORM2 plants showed a dramatic increase in the total sphingolipid contents, especially in LCBs (∼15-fold) and ceramides (8-fold), compared with the wild type. By contrast, the amount of glucosylceramides did not change relative to the wild type (Figure 1E). Among LCBs, we noticed that d18:0 LCB showed the largest difference in orm1 amiR-ORM2 (Supplemental Figure 4). Remarkably, the ceramide backbones containing long-chain fatty acids (C16) exhibited a more dramatic increase than ceramide containing very-long-chain fatty acids (C20, C24, and C26) (Supplemental Figure 4).

We also measured the sphingolipid contents of orm1 amiR-ORM2 and wild-type rosettes at different developmental stages, and the results showed a gradual accumulation of sphingolipids in orm1 amiR-ORM2 plants (Supplemental Figure 5). The sphingolipid profile of 21-d-old plants revealed that the total amounts of LCBs, ceramides, and hydroxyceramides were higher in orm1 amiR-ORM2 plants compared with wild-type plants. However, no visible early-senescence phenotype was observed in the orm1 amiR-ORM2 plants at this stage (Supplemental Figure 3). In other words, orm1 amiR-ORM2 plants accumulated sphingolipids prior to the early senescence. There were no significant changes in sphingolipids in orm1 mutants, amiR-ORM2 plants, or overexpression transgenic plants with respect to wild-type plants at different developmental stages (Supplemental Figures 3 and 5). These results indicated that ORM1 and ORM2 negatively coregulate sphingolipid biosynthesis in plants. In addition, the early-senescence phenotype of orm1 amiR-ORM2 plants was associated with overaccumulation of sphingolipids.

ORMs Suppress the de Novo Biosynthesis of Sphingolipids in Arabidopsis

To decipher the possible cause of the overaccumulation of sphingolipids, we used 15N-labeling and metabolic turnover analysis to directly measure in vivo sphingolipid changes, as we reported previously (Shi et al., 2015). Seven-day-old orm1 amiR-ORM2 and wild-type seedlings were transferred to N-deficient 0.5× Murashige and Skoog (MS) liquid medium containing 5 mM 15N-serine in a time course. Isotope-labeled LCBs and ceramides of orm1 amiR-ORM2 plants accumulated to significant levels after 24 h of incubation with 15N-serine, relative to wild-type and orm1 or amiR-ORM2 plants (Figure 2). It is noteworthy that the increase of 15N-labeled d18:0 LCB was greater than that of t18:0 LCB (Figure 2). In addition to quantitative changes in ceramides, d18:0 ceramides showed a more dramatic increase than t18:0 ceramides (Figure 2). The initial LCB produced in plants is dihydrosphingosine (d18:0 LCB) (Chen et al., 2009). Hence, these measurements confirmed our hypothesis that the loss of ORM1 and ORM2 function in the orm1 amiR-ORM2 plants promotes the de novo biosynthesis of sphingolipids and ultimately leads to higher accumulation of sphingolipids. Moreover, the higher levels of 15N-labeled ceramides containing C16 fatty acid in orm1 amiR-ORM2 plants (Figure 2) were consistent with accumulation of C16 ceramides in 28-d-old orm1 amiR-ORM2 plants (Supplemental Figure 4).

Figure 2.

Figure 2.

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ORM Suppresses the de Novo Biosynthesis of Sphingolipids in Arabidopsis.

Seven-day-old wild-type, orm1, amiR-ORM2, and orm1 amiR-ORM2 seedlings were transferred to 5 mM 15N-serine labeled N-deficient 0.5× MS liquid medium for 9 to 24 h. Sphingolipids were then extracted and measured as described in Methods. Note that except total LCB and total ceramides (top panels), major LCB and ceramide species significantly increased at 24 h in orm1 amiR-ORM2 seedlings. Error bars represent the means ± se from triplicate biological repeats. Asterisks show a significant difference from the wild type using Student’s t test (*P < 0.05, **P < 0.01, and ***P < 0.001).

ORM1 and ORM2 Interact with ssSPT

Previous studies in yeast and mammals showed that ORM proteins physically interact with Lcb1 and Lcb2 (Han et al., 2010). To test the interactions of ORM proteins in Arabidopsis, we first applied yeast two-hybrid assays. Surprisingly, our yeast two-hybrid assays revealed that the ORM proteins physically interact with the ssSPT (Figure 3A). The interaction was further confirmed by coimmunoprecipitation (co-IP) assays. Anti-Flag resins could precipitate not only ORM-Flag but also GFP-ssSPT (Figure 3B). When free GFP was used as a control, no GFP signals were detected in the eluate (Figure 3B). We also used bimolecular fluorescence complementation (BiFC) assays to confirm the ORM-ssSPT interaction. Clear YFP fluorescence was observed in protoplasts cotransformed with pSATN-nEYFP-ORM1 and pSATN-cEYFP-ssSPTa or pSATN-cEYFP-ssSPTb, pSATN-nEYFP-ORM2, and pSATN-cEYFP-ssSPTa or pSATN-cEYFP-ssSPTb constructs (Figure 3C). However, no YFP signal was detected in the protoplasts cotransformed with one construct in combination with an empty vector (Figure 3C). In addition, our confocal microscopy observations confirmed that ssSTPa and ssSTPb also localized to the ER where they interact with ORMs (Supplemental Figure 6). These results are consistent with the physical interaction of ORM1 and ORM2 with ssSPTa and ssSPTb at the ER. We also found that SPTs localized to ER in the orm1 amiR-ORM2 plants, as confirmed by transient coexpression with 35Spro:EGFP-LCB1 or 35Spro:EGFP-LCB2a/b and an ER marker, suggesting that loss of function of ORMs did not affect subcellular localization of the SPT complex (Supplemental Figure 7).

Figure 3.

Figure 3.

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Physical Interaction between ORM and ssSPT.

(A) Yeast two-hybrid assay showing that ssSPTa and ssSPTb interact with ORM1 and ORM2. Vectors pGADT7 (AD) and pGBKT7 (BD) were used as negative controls.

(B) Co-IP assays of ORM1, ORM2, ssSPTa, and ssSPTb in protoplasts. Flag-tagged ORM1 and ORM2 were immunoprecipitated with an anti-Flag antibody, and coimmunoprecipitated ssSPTa-GFP and ssSPTb-GFP were detected by an anti-GFP antibody. The “+” or “−”denote the presence or absence of the protein in each sample. Free GFP was used as a negative control.

(C) BiFC interaction of ssSPTa and ORM1, ssSPTb and ORM2, ssSPTa and ORM2, and ssSPTb and ORM2 in Arabidopsis leaf protoplasts. Overlaid images show signals for YFP (yellow) and chloroplasts (blue). Empty vectors were used as negative controls. Bars = 5 μm.

ORM Proteins Decrease the Biosynthesis of Sphingolipids

Reduced SPT activity may decrease sensitivity to fumonisin B1 (FB1) by decreasing levels of cytotoxic LCBs, whereas increased SPT activity may increase sensitivity to FB1 by increasing the levels of LCBs (Shi et al., 2007; Saucedo-García et al., 2011). ssSPTa/ssSPTb can strongly stimulate SPT activity and increase the production of LCBs in Arabidopsis (Kimberlin et al., 2013). To explore the function of Arabidopsis ORMs, we grew plants on agar plates with FB1 to test the significance of the interaction between ORM and ssSPT. Both ORM1-OX and ORM2-OX overexpression lines showed resistance to 0.5 μM FB1, whereas the orm1 amiR-ORM2 plants were highly sensitive to FB1 (Figure 4). Consistent with a previous human ORM study (Breslow et al., 2010), ORM proteins thus appear to negatively regulate sphingolipid biosynthesis.

Figure 4.

Figure 4.

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Phenotypes of Wild-Type and ORM Transgenic Seedlings after FB1 Treatments.

Seedlings of wild-type, orm1, amiR-ORM2, and orm1 amiR-ORM2 plants and overexpression lines (ORM1-OX and ORM2-OX) were grown on 0.5× MS with or without 0.5 μM FB1 for 9 d and then photographed. Percentage of seedling mortality after 0.5 μM FB1 treatment is shown in the right panel. Asterisks show a significant difference from the wild type using Student’s t test (*P < 0.05, **P < 0.01, and ***P < 0.001).

Loss of ORM Proteins Increases ER Stress-Related Responses

To understand whether the loss of ORM proteins affects cellular ultrastructure, we fixed the orm1 amiR-ORM2 leaves before the appearance of the senescence phenotype. As shown in Figure 5, compared with the normal nuclei observed in the wild-type cells (Figure 5A), the orm1 amiR-ORM2 cells showed condensed chromatin aggregated in the perinuclear membrane, which indicates dying cells (Figure 5B; Liang et al., 2003), and a large amount of tiny membrane sacs close to the plasma membrane and the cell wall (Figures 5C and 5D). Interestingly, we found a large cell wall apposition, a clear marker of the defense response, in the orm1 amiR-ORM2 leaf cells (Figure 5E). When we used the cerium chloride method (Bi et al., 2014) to detect reactive oxygen species (ROS), we observed H2O2 production on the cell wall (Figures 5G and 5H), the cytosolic area close to the apoplast (Figure 5I) and in the mitochondria (Figure 5J). No cerium deposits were observed in the wild-type control (Figure 5F). In the dying cells, irregular ER and vacuolization were frequently observed (Figure 5K). These observations suggest that loss of ORMs may induce active vesicular trafficking around the plasma membrane, and this ER stress phenomenon is associated with defense responses.

Figure 5.

Figure 5.

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Ultrastructural Features of orm1 amiR-ORM2 Leaves.

(A) to (E) Representative TEM images of ultrastructure of 25-d-old wild-type (A) and orm1 amiR-ORM2 ([B] to [E]) leaves. Note an aggregate of condensed chromatin ([B], white arrow), abnormal ER with loose structure (C), and many small bubbles ([C] and [D], white stars) around the plasma membrane and cell wall and a large cell wall apposition in the orm1 amiR-ORM2 leaf cell (E).

(F) to (K) H2O2 production in wild-type (F) and orm1 amiR-ORM2 ([G] to [K]) leaves observed by TEM using the histochemical cerium chloride method.

(F) No cerium deposits were observed in the representative wild-type control cell.

(G) and (H) Cerium deposits in the plasma membrane and cell wall (black arrows). Note vesicular bodies with double membrane (black arrowheads).

(I) Cerium deposits around the ER close to the cell wall.

(J) and (K) H2O2 inside mitochondria ([J], white arrowheads) and irregular vacuolization (K).

Ch, chloroplast; CW, cell wall; CWA, cell wall apposition; G, Golgi; M, mitochondrion; N, nucleus; S, starch grain. Bars = 500 nm.

To support the transmission electron microscopy (TEM) observations, we investigated the expression of genes related to ER stress and defenses. Strikingly, the expression of ER stress marker genes (bZIP28, bZIP60, IRE1a, and TBF1) significantly increased in orm1 amiR-ORM2 plants, compared with the wild type (Figure 6A). To determine whether the transcription of defense-related genes was affected in plants with decreased ORM function, we analyzed the expression of salicylic acid-related and pathogenesis-related (PR) genes in wild-type and orm1 amiR-ORM2 plants using RT-qPCR. The orm1 amiR-ORM2 plants showed high transcript levels of these genes, such as PAD4, SID2, and NPR1, compared with the wild type (Figure 6B). PR1 showed especially dramatic enhancement, which is highly linked to the formation of cell wall appositions. No obvious changes of those genes were detected in orm1 and amiR-ORM2 plants (Figure 6).

Figure 6.

Figure 6.

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Expression of Genes Related to ER Stress and Defense in orm1 amiR-ORM2 Plants.

Approximately 26-d-old wild-type, orm1, amiR-ORM2, and orm1 amiR-ORM2 plants were used to monitor the expression of indicated genes by RT-qPCR. IRE1a (At2G17520), bZIP28 (At3G10800), bZIP60 (At1G42990), TBF1 (At4G36990), PR1 (At2g14610), PAD4 (At3g52430), SID2 (At1g74710), and NPR1 (At1g64280) were used for analysis. ACT2 transcript levels were used as the internal control. ER stress-related genes (A) and salicylic acid pathway-related genes (B). Gene expression values are presented relative to average wild-type levels (set to 1). Samples were analyzed in triplicate biological repeats with similar results. Bars indicate ± se from three technical replicates. Different letters represent a significant difference from the wild type using a post hoc multiple t test (P < 0.001).

Loss of ORM Proteins Enhances Plant Resistance to Abiotic and Biotic Responses

Recent studies have shown that sphingolipids are involved in plant stress responses (Wang et al., 2008; Peer et al., 2010; Bi et al., 2014; Li et al., 2015). The orm1 amiR-ORM2 plants exhibit highly induced expression of ER stress and salicylic acid-related genes, which suggested to us that we should explore the role of ORMs during abiotic and biotic stress. We first treated the orm1 amiR-ORM2 plants with the oxidative stress agent methyl viologen (MV), which produces ROS. Surprisingly, orm1 amiR-ORM2 plants proved to be tolerant to ROS and exhibited dramatically higher survival rates than wild type (Figures 7A and 7B).

Figure 7.

Figure 7.

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Loss of ORM Protein Enhances the Resistance to Abiotic Stress and Biotic Stress.

(A) The phenotype of wild-type and mutant seedlings after MV treatments. Seedlings were grown on 0.5× MS with or without 0.5 μM MV for 7 d and then photographed. The white square indicates one seedling with a severe phenotype.

(B) Percentage of seedlings with a severe phenotype after 0.5 μM MV treatments. At least 350 seedlings were counted in each genotype. Values represent means ± se from two independent experiments.

(C) Cell death symptom after infection by P. syringae virulent strain DG3 at OD600 = 0.005 for 3 d. MgSO4 as a mock-inoculated control was used.

(D) Growth of P. syringae virulent strain DG3 in wild-type and orm1 amiR-ORM2 plants after infection at OD600 = 0.001. Lesion-free plants were inoculated with bacteria at 3 weeks of age. The mean value of the growth of bacteria in nine leaves is indicated in each case. Bars indicate standard deviations. This experiment was repeated three times with similar results. Asterisks show a significant difference from the wild type using Student’s t test (*P < 0.05).

We further inoculated orm1 amiR-ORM2 plants with the bacterial pathogen Pseudomonas syringae strain DG3. The orm1 amiR-ORM2 plants showed subtle symptoms after infection and proved to be relatively resistant to DG3 bacteria when compared with the wild type (Figure 7C). This enhanced disease resistance was consistent with the detection of high transcript levels of resistance genes in orm1 amiR-ORM2 plants. These results indicate that loss of ORM function in Arabidopsis affects the plant’s response to abiotic and biotic stress.

DISCUSSION

The ORM proteins, encoded by ORMDL (ORM) genes, are highly conserved eukaryotic transmembrane proteins located in the ER (Hjelmqvist et al., 2002; Han et al., 2010). The three ORMDL proteins are highly conserved in humans and yeast has two homologous ORM genes (ORM1 and ORM2) with redundant functions. Compared with other organisms, the extent of conservation of ORM function in plants remains unclear. In this report, we characterized the two ORM genes found in Arabidopsis. Our results indicate that Arabidopsis ORMs function to regulate sphingolipid biosynthesis in maintaining sphingolipid homeostasis and play a role in response to abiotic and biotic stresses.

ORM proteins have few conserved regions, mainly located in the middle of the amino acid sequence, and are generally predicted to form two to four transmembrane domains. The amino acid sequences of the Arabidopsis ORMs share 81% identity and also share 35 to 39% identity with S. cerevisiae Orm1 and Orm2 (Supplemental Figure 1), but interestingly, the Arabidopsis isoforms are N-terminally truncated relative to the yeast proteins and therefore lack the three serine residues that are phosphorylated by Ypk1 in the yeast proteins (Roelants et al., 2011). Although we found some possible phosphorylation sites in Arabidopsis ORM1 and ORM2, we still do not know whether the ORMs in Arabidopsis are regulated by phosphorylation, similar to yeast Orm proteins. Another possibility is that Arabidopsis ORMs may be regulated by an allosteric effect on the ORM proteins themselves, where sphingolipids trigger a change in conformation within a preexisting ORM/SPT complex rather than enhancing formation of the complex (Kiefer et al., 2015).

Orm1 and 2 negatively regulate SPT, activating or inhibiting its activity through phosphorylation and dephosphorylation (Tafesse and Holthuis, 2010). Regulation of sphingolipids and Orm proteins involves a feedback mechanism; Orms regulate SPT and sphingolipid metabolites such as LCBs, ceramide, and accumulated sphingoid intermediates, and then trigger Orm dephosphorylation, which in turn downregulates sphingolipid biosynthesis (Sun et al., 2012). In yeast, Orm1 and Orm2 are mainly phosphorylated by Npr1 and Ypk1, respectively (Roelants et al., 2011; Sun et al., 2012; Shimobayashi et al., 2013). Besides ORMs, other polypeptides can also regulate the activity of SPT, namely, Tsc3 in yeast (Gable et al., 2000) and ssSPTa (68 amino acids) and ssSPTb (76 amino acids) in human (Han et al., 2009). The 56-amino acid ssSPTs, ssSPTa and ssSPTb, strongly stimulate Arabidopsis SPT activity when coexpressed with Arabidopsis LCB1 and LCB2a or 2b in a yeast spt null mutant (Kimberlin et al., 2013). Also, ssSPTa suppression lines increase the resistance to FB1, and the ssSPTa overexpression lines display strong sensitivity to FB1 by increasing or decreasing SPT activity (Kimberlin et al., 2013). In this study, the de novo biosynthesis of sphingolipids was activated due to loss of Arabidopsis ORM function. The isotope-labeled LCBs and ceramides of orm1 amiR-ORM2 accumulated significantly after 24 h of incubation in 15N-labeled serine, relative to the wild type, which showed that ORM was indeed a negative regulator of SPT in Arabidopsis, like in human and yeast. Through a yeast two-hybrid system, BiFC experiments, and co-IP assays, we confirmed that Arabidopsis ORMs can directly bind with the Arabidopsis ssSPTs. Based on our data, we put forward two possible mechanisms for the function of ORMs (Figure 8). On the one hand, ORM protein might bind to the SPT complex through the physical interaction with ssSPT and inhibit the enzyme activity of SPT, rather than changing its localization; on the other hand, the interaction between ORM and ssSPT might reduce the available ssSPTs that can interact with core LCB1/LCB2. We favor the first mechanism, in which ORM binds to the SPT complex with the ssSPT interaction, based on a recent study by Kimberlin et al. (2016). In addition, we observed that the ORMs and ssSPT localize in the ER, which could be the site where they interact (Figure 1; Supplemental Figure 6). Moreover, during FB1 treatment, ORM overexpression lines displayed strong resistance to FB1, the opposite effect to that of ssSPTa. Taking these results together, we speculate that ORMs could modulate sphingolipid levels by inhibiting the function of ssSPTs in Arabidopsis. When the ORMs are knocked out, SPT could be activated without changing its location, resulting in substantial accumulation of sphingolipids (Figure 8).

Figure 8.

Figure 8.

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Model of ORM1- and ORM2-Mediated Sphingolipid Homeostasis and Plant Resistance.

The proposed model for ORM1 and ORM2 functions as negative regulators of de novo sphingolipid synthesis in Arabidopsis based on this study. SPT, localized on the ER, consists of LCB1, LCB2a/b, and ssSPTa/b and catalyzes the first step in sphingolipid biosynthesis. SPT demonstrates extremely low enzyme activity without ssSPTa/b (Kimberlin et al., 2013). We found that ORM1 and ORM2 inhibit the biosynthesis of LCB through physically interacting with ssSPTa/b. Moreover, in plants lacking ORM1 or ORM2, the profile of sphingolipids was enhanced, accompanied by a senescence phenotype and increased disease resistance. A possible mechanism related to defense in ORM1 and ORM2 loss-of-function plants is proposed. CW, cell wall; SA, salicylic acid; CWA, cell wall apposition. Red letters and arrows represent data in this study. The black dotted arrow represents our speculation.

Like in humans and yeast, ORM proteins in Arabidopsis play an important role in maintaining the level of sphingolipids (Tafesse and Holthuis, 2010; Breslow et al., 2010; Liu et al., 2012). In this study, we analyzed the ORM1 T-DNA insertion mutant orm1 and the ORM2 silencing line (amiR-ORM2). Under normal growth conditions, these plants showed no obvious differences compared with the wild type. However, the orm1 amiR-ORM2 plants exhibited premature senescence and considerable changes in sphingolipid contents, including multiple-fold increases in overall sphingolipids, especially in LCBs and ceramides. Among these, d18:0 and t18:0 LCB, the ceramide backbones containing long-chain fatty acids (C16), exhibited the largest changes. Measurements of the sphingolipid contents of orm1 amiR-ORM2 and wild-type rosettes at different developmental stages showed that prior to the onset of the early-senescence phenotype, the total LCBs, ceramides, and hydroxyceramides, mainly ceramide backbones containing dihydroxy LCBs and C16 fatty acids, increased in orm1 amiR-ORM2 plants compared with wild-type plants. These observations suggest that LCBs or ceramides may induce plant senescence and accumulate to threshold levels due to loss of ORM function.

Sphingolipids are involved in the regulation of plant responses to biotic and abiotic stresses. Our previous studies showed that the Arabidopsis neutral ceramidase mutant ncer1 accumulates hydroxyceramides and is sensitive to oxidative stress (Li et al., 2015) and the ceramide kinase mutant acd5 accumulates ceramides and is sensitive to P. syringae and Botrytis cinerea (Liang et al., 2003; Bi et al., 2014). In addition, the fatty acid α hydroxylase mutant fah1 fah2 has a stronger resistance to Diplodia powdery mildew and Verticillium fungi (Verticillium longisporum) compared with the wild type (König et al., 2012). In this study, we found that plants with loss of ORM function are more resistant to MV and pathogen infection. This phenomenon could be attributed to the higher accumulation of LCBs in the ORMs mutants compared with the acd5 and fah1 fah2 mutants. Several previous studies reported that LCBs play an important role during abiotic and biotic stress (Peer et al., 2010; Saucedo-García et al., 2011; Li et al., 2015). LCBs can induce MPK6 expression and programmed cell death by an MPK6-mediated signal transduction pathway (Saucedo-García et al., 2011). The expression of MPK6 can be induced rapidly by Flg22, which mediates the pathogen-associated pattern-induced basal resistance (Galletti et al., 2011). Phytosphingosine content has been reported to rapidly escalate at two hours after P. syringae inoculation and is also involved in plant resistance (Peer et al., 2010). Our sphingolipid data also showed an increase in the phytosphingosine contents in plants with loss of ORM function, which leads to increased resistance against P. syringae infection. In addition, our TEM observation gives us another indication of the defense response in plants with decreased ORM1 or ORM2 (Figure 8), showing that active vesicular transport may be caused by perturbing the ORM-mediated sphingolipid homeostasis, and this may strengthen cell surface defenses (including formation of cell wall appositions and ROS production on the cell wall). Furthermore, based on the high transcript levels of resistance-related genes (PR1) in plants with loss of ORM function, and especially the upregulated ER stress-related genes (Liu and Howell, 2010) and expression of PAD4, SID2, and NPR1 genes related to the SA pathway, we propose that loss of ORM function may affect SA biosynthesis, thereby affecting the plant’s resistance to pathogens.

METHODS

Plant Materials and Growth Conditions

Wild-type Arabidopsis thaliana (Columbia, Col-0) and T-DNA insertion orm mutants (SALK_046054 and SAIL_1286_D09) from the ABRC (http://abrc.osu.edu/) were sown on soil after 3 d of stratification at 4°C, followed by cultivation in the greenhouse at 22°C and 50% relative humidity, 16 h light/8 h dark with 4800 to 6000 lux light intensity (PAK bulb, PAK090311).

To construct the ORM1 and ORM2 RNAi suppression vectors, the RS300 plasmid was employed to clone an artificial microRNA, as previously described (Schwab et al., 2006). The artificial microRNA was inserted into pCAMBIA1300 and fused with the 35S promoter and NOS terminator. The ORM1 and ORM2 overexpression constructs were generated using pCAMBIA1300 by inserting the open reading frame of ORM1 or ORM2, the 35S promoter, and NOS terminator sequences. Finally, these constructs were transformed into the wild type using an Agrobacterium tumefaciens (EHA105)-mediated method. Transformed progenies were screened on 0.5× MS medium containing 0.25 mg/L hygromycin. Homozygous transgenic lines were isolated from the T3 generation for further study. All primers used for cloning are shown in Supplemental Table 1.

Phosphorylation Site Analysis of ORMs

For the analysis of potential phosphorylation sites presented in ORM proteins, the amino acid sequences of Arabidopsis ORM1 and ORM2 were submitted to the NetPhos 3.1 (http://www.cbs.dtu.dk/services/NetPhos/) and PhosPhAt 4.0 (http://phosphat.uni-hohenheim.de/phosphat.html) online tools, respectively (Blom et al., 2004; Durek et al., 2010).

Quantitative RT-PCR Analysis

Total RNA was extracted using the E.Z.N.A. plant RNA kit (R6827-01; Omega Bio-tek). For each sample, 1 μg RNA was reverse transcribed into cDNA using the Primescript RT reagent kit (Takara; DRR047A). Real-time PCR was performed with the SYBR Premix ExTaq kit (Takara; RR820L) according to the manufacturer’s instructions and quantitatively analyzed with a Step One Plus real-time PCR system (ABI). The 2−△△CT method (Livak and Schmittgen, 2001) was used to determine the relative transcript level of target genes according to the expression level of ACT2 (the internal control). All the experiments were performed in triplicate. The primers used in this study are listed in Supplemental Table 1. Unless otherwise mentioned, all chemicals were purchased from Sigma-Aldrich.

Subcellular Protein Localization

For subcellular protein localization, the 35Spro:GFP:ORM1 and 35Spro:GFP:ORM2 gene expression cassettes were constructed and inserted into pCAMBIA1300. Mesophyll protoplasts were isolated by the tape-Arabidopsis sandwich method and transformed by PEG-calcium mediated transfection (Wu et al., 2009). The transfected protoplasts were cultured under dim light (∼300 lux) for 16 to 24 h at room temperature and observed by confocal microscopy (LSM-780; Carl Zeiss). The excitation/emission wavelengths were: 488 nm/500 to 530 nm for GFP, 561 nm/580 to 630 nm for mCherry, and 488 nm/650 to 750 nm for chlorophyll.

Sphingolipid Analysis

Measurement of sphingolipids was performed and the data were analyzed by a Shimadzu UFLC-XR coupled with a hybrid quadrupole time-of-flight mass spectrometer (AB SCIEX Triple TOF 5600+) using a Phenomenex Luna C8 column (150 mm × 2.0 mm, 3 μm). Briefly, 30 mg of lyophilized sample was homogenized. The internal standards (C17 base d-erythro-sphingosine and d18:1 C12:0-ceramide) were added and extracted with the isopropanol/hexane/water (55:20:25 v/v/v) and incubated at 60°C for 15 min. After centrifugation, the supernatants were dried and de-esterified in methylamine in ethanol/water (70:30 v/v) as described previously (Bi et al., 2014; Li et al., 2015; Wu et al., 2015). The sphingolipid species were analyzed using the software Multiquant (AB SCIEX).

Yeast Two-Hybrid Assay

Yeast two-hybrid analysis was conducted following the Matchmaker Gold Yeast Two-Hybrid System User Manual (Clontech). The full-length open reading frames of ssSPTa and ssSPTb were fused to the bait vector pGBKT7, and the full-length open reading frames of ORM1 and ORM2 were cloned into the prey vector pGADT7. Prey and bait vectors were transformed into the yeast strain Y2H Gold (Clontech), and yeast was grown on SD/-Trp-Leu medium for 3 d. Transformants were incubated at 30°C in a shaking incubator with SD/-Trp-Leu broth until OD600 = 1.0 was obtained and then tested on selective SD medium at 30°C for 5 d. Empty vectors were used as the negative control.

BiFC Assay

The full-length coding sequences of ORM1 and ORM2 were cloned into pSATN-nEYFP-C1, and full-length coding sequences of ssSPTa and ssSPTb were cloned into pSATN-cYFP-C1 (Citovsky et al., 2006). Protoplast isolation and transient expression were performed as described previously (Wu et al., 2009). Empty vectors were cotransformed as negative controls.

Co-IP Assay

Mesophyll protoplast isolation from 3- to 4-week-old Arabidopsis leaves and DNA transfection were performed as described previously (Wu et al., 2009). For the protocol, 100 μg of prey plasmids and 100 μg of bait plasmids were cotransfected into 1 mL protoplasts (5 × 105 cells). After expression of proteins for 12 h, protoplasts were pelleted and lysed in 200 μL of immunoprecipitation buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM EDTA, 10% glycerol, 1% Triton X-100, and 13 Roche EDTA-free protease inhibitor cocktail) by vigorous vortexing. For each sample, 20 μL of lysate was saved as the input fraction. The remaining of the lysate was mixed with 300 μL immunoprecipitation buffer and vigorously vortexed. The clear lysate was centrifuged at 16,000g for 10 min at 4°C, and the supernatant was incubated with 20 μL of anti-Flag agarose resins (Sigma-Aldrich) for 4 h at 4°C. The resin was washed three times with immunoprecipitation buffer. The resins were boiled in 40 μL of SDS-PAGE loading buffer to obtain the eluate and prey proteins was detected by immunoblotting analysis using anti-Flag antibody and anti-GFP antibody (Cell Signal) at 1:2000 dilution. The immunoblot signal was visualized with the Clarity Western ECL substrate kit (Bio-Rad).

H2O2 Detection by CeCl3 Staining and TEM Observation

For electron microscopy samples, ∼25-d-old Arabidopsis rosette leaves from soil-grown plants were used. The histochemical cerium chloride method was used to detect H2O2 based on generation of cerium hydroxide, as described previously (Bi et al., 2014). The leaves were cut and incubated in 10 mM CeCl3 dissolved in 50 mM MOPS buffer (pH 7.2) for 1 h. Control samples were incubated in MOPS buffer only. Samples were fixed in 2.5% (v/v) glutaraldehyde and 2% (v/v) paraformaldehyde in 0.1 M cacodylate buffer (pH 7.4). Samples were embedded in SPI-PON812 resin (SPI Supplies). Ultrathin sections were obtained on a microtome (Leica EM UC6) and examined without staining. The images were photographed using a transmission electron microscope (JEM-1400; JEOL) at an accelerating voltage of 120 kV.

Abiotic and Biotic Stress Treatments

The seeds of each line were sown on 0.5× MS medium supplemented with FB1 (0.5 μM) or MV (0.5 μM). After 2 to 3 d at 4°C in the dark, plates were transferred into an incubator with a 16-h-light/8-h-dark light regimen. Phenotypes were characterized and scored. For bacterial infection, leaves from 3-week-old plants were injected with the virulent Pseudomonas syringae strain DG3 at OD600 = 0.001 or with 10 mM MgSO4 as a mock-inoculated control. Leaf discs were harvested for bacterial quantification at indicated days after inoculation as previous reports (Bi et al., 2014; Wu et al., 2015).

Accession Numbers

Sequence data from this article can be found in the Arabidopsis Genome Initiative or GenBank/EMBL databases under the following accession numbers: ORM1 (At1G01230), ORM2 (At5G42000), ssSPTa (At1g06515), ssSPTb (At2g30942), LCB1 (At4g36480), LCB2a (At5g23670), LCB2b (At3g48780), ACT2 (At3g18780), TUBULIN (At5G62690), IRE1a (At2G17520), bZIP28 (At3G10800), bZIP60 (At1G42990), TBF1 (At4G36990), PAD4 (At3g52430), SID2 (At1g74710), NPR1 (At1g64280), PR1 (At2g14610), orm1 (SALK_046054), and orm2 (SAIL_1286_D09).

Supplemental Data

Supplementary Material

Supplemental Data
supp_28_12_3038__index.html (1.2KB, html)

Acknowledgments

We thank Edgar Cahoon (University of Nebraska-Lincoln) for helpful discussion and Jian-Feng Li (Sun Yat-sen University) for assistance with co-IP experiments. This work was funded by the National Natural Science Foundation of China (31570255), the National Key Basic Science 973 program (2012CB114006), the Foundation of Guangzhou Science and Technology Project (201504010021), and the Fundamental Research Funds for the Central Universities (16lgjc74) to N.Y. and by the China Postdoctoral Science Foundation (2016M592575) to S.K.S.

AUTHOR CONTRIBUTIONS

N.Y., J.L., and J.-X.W. conceived and designed experiments. J.L., J.Y., C.R., K.-E.L., L.-Q.H., and H.-Y.Z. performed the experiments. J.L., J.Y., K.-E.L., J.-X.W., and N.Y. analyzed the data. N.Y, J.L., and S.K.S. wrote the article.

Glossary

SPT

serine palmitoyltransferase

ER

endoplasmic reticulum

ssSPT

SPT small subunit

LCB

long-chain base

MS

Murashige and Skoog

co-IP

coimmunoprecipitation

BiFC

bimolecular fluorescence complementation

ROS

reactive oxygen species

MV

methyl viologen

TEM

transmission electron microscopy

Footnotes

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