Skip to main content
PMC home page
The Plant Cell logoLink to The Plant Cell
. 2011 May 24;23(5):1971–1984. doi: 10.1105/tpc.110.081943

The Plant Cuticle Is Required for Osmotic Stress Regulation of Abscisic Acid Biosynthesis and Osmotic Stress Tolerance in Arabidopsis[W]

Zhen-Yu Wang a,b, Liming Xiong a, Wenbo Li c, Jian-Kang Zhu a,c, Jianhua Zhu d,1
PMCID: PMC3123942  PMID: 21610183

The plant cuticle is well known as a physical cover for the plant’s aerial parts, preventing water loss and invasion by pathogens and pests. This study found a surprising role of cutin, a major component of the cuticle, in osmotic stress induction of biosynthesis of the stress hormone abscisic acid.

Abstract

Osmotic stress activates the biosynthesis of abscisic acid (ABA). One major step in ABA biosynthesis is the carotenoid cleavage catalyzed by a 9-cis epoxycarotenoid dioxygenase (NCED). To understand the mechanism for osmotic stress activation of ABA biosynthesis, we screened for Arabidopsis thaliana mutants that failed to induce the NCED3 gene expression in response to osmotic stress treatments. The ced1 (for 9-cis epoxycarotenoid dioxygenase defective 1) mutant isolated in this study showed markedly reduced expression of NCED3 in response to osmotic stress (polyethylene glycol) treatments compared with the wild type. Other ABA biosynthesis genes are also greatly reduced in ced1 under osmotic stress. ced1 mutant plants are very sensitive to even mild osmotic stress. Map-based cloning revealed unexpectedly that CED1 encodes a putative α/β hydrolase domain-containing protein and is allelic to the BODYGUARD gene that was recently shown to be essential for cuticle biogenesis. Further studies discovered that other cutin biosynthesis mutants are also impaired in osmotic stress induction of ABA biosynthesis genes and are sensitive to osmotic stress. Our work demonstrates that the cuticle functions not merely as a physical barrier to minimize water loss but also mediates osmotic stress signaling and tolerance by regulating ABA biosynthesis and signaling.

INTRODUCTION

The phytohormone abscisic acid (ABA) regulates many aspects of plant growth and development as well as responses to the environment (Borsani et al., 2002; Finkelstein et al., 2002; Zhu, 2002; Nambara and Marion-Poll, 2005). Plants maintain a low level of ABA under normal growth conditions. This level of ABA may be required for normal plant growth and development since in strong ABA-deficient mutant backgrounds, or when ABA signaling is blocked, plants have difficulty surviving even under relatively normal growth conditions (Fujii and Zhu, 2009). In response to environmental stress, such as drought or high salinity, ABA levels increase dramatically to cope with the stress. This stress regulation of ABA biosynthesis primarily occurs at the transcriptional level, although other levels of control, such as protein stability or activity, conjugation, and catabolism, may also play important roles in regulating ABA levels (Finkelstein et al., 2002; Xiong and Zhu, 2003; Okamoto et al., 2009; Cutler et al., 2010).

De novo ABA biosynthesis in higher plants proceeds through the cleavage of a C40 carotenoid precursor, followed by a two-step conversion of the intermediate xanthoxin to ABA via ABA-aldehyde (Taylor et al., 2000; Seo and Koshiba, 2002; Schwartz et al., 2003; Nambara and Marion-Poll, 2005; Finkelstein and Rock, 2008). All these steps in the ABA biosynthesis pathway may be regulated to various extents, as it was discovered that ABA biosynthesis genes are upregulated both by stress and by ABA (Xiong et al., 2001, 2002; Xiong and Zhu, 2003). The stress regulation and self-regulation of the pathway may thus provide plants a mechanism for rapid response to environmental challenges. Although ABA biosynthesis may be regulated at multiple steps, the cleavage step catalyzed by a 9-cis epoxycarotenoid dioxygenase (NCED) is generally considered to be the rate-limiting step (Tan et al., 1997; Qin and Zeevaart, 1999; Iuchi et al., 2001). Thus, determining how NCED genes are regulated by osmotic stress will be instrumental for understanding the mechanisms of plant acclimation to stress.

To begin to dissect the regulatory circuits for stress induction of ABA biosynthesis, we adopted a genetic approach using an Arabidopsis NCED3 promoter driven firefly luciferase (LUC) reporter system to isolate mutants defective in stress regulation of ABA biosynthesis. Since osmotic stress is an important factor for drought stress, and osmotic stress treatment could be administered in a more quantitative and reproducible way using polyethylene glycol (PEG) in agar media, we chose to investigate osmotic stress induction of the NCED3:LUC reporter gene. In this study, we identified and characterized a regulator of NCED3 expression, CED1 (for 9-cis epoxycarotenoid dioxygenase defective 1). In ced1 mutant plants, the induction of NCED3 by osmotic stress was greatly reduced. The mutant plants also had a lower ABA level and were extremely sensitive to osmotic stress. Unexpectedly, CED1 was found to encode a putative α/β hydrolase domain-containing protein and is allelic to BODYGUARD (BDG), which has been shown to be essential for cutin synthesis and disease resistance. Although it has long been known that cuticle plays important roles in minimizing water loss and increasing plant resistance to both biotic and abiotic challenges, the discovery of BDG/CED1 as important for osmotic stress induction of ABA biosynthesis and osmotic stress tolerance suggests that this thin physical barrier is actively involved in signaling of not only biotic but also abiotic stresses to enhance plant survival under adverse environmental conditions.

RESULTS

ced1 Mutant Plants Are Impaired in Osmotic Stress–Induced ABA Accumulation

To understand the mechanisms of stress regulation of ABA biosynthesis, we took a genetic approach to analyze stress induction of the NCED3 gene. Transgenic Arabidopsis thaliana plants expressing the firefly LUC reporter gene under control of the NCED3 promoter (NCED3:LUC) (referred as the wild type hereafter) were generated, and the resulting seeds were mutagenized using ethyl methanesulfonate. We initially screened and obtained a large number of mutant lines with an altered regulation of NCED3:LUC in response to osmotic stress. In subsequent generations, however, the expression of the LUC reporter gene in these mutants and the wild type was silenced probably due to increased DNA methylation in the NCED3 promoter region of the transgene since treatment with the DNA methylation inhibitor 5-Aza-C could restore NCED3:LUC expression (see Supplemental Figure 1 online). With the expectation that changes in NCED3 expression may affect ABA biosynthesis and stress resistance, we rescreened these putative mutant lines for altered ABA accumulation and resistance to low water potential stress. This resulted in the identification of a group of mutants that show lower levels of ABA accumulation compared with the wild type when treated with osmotic stress. One such mutant, ced1, was chosen for further characterization in this study.

ced1 mutant plants were backcrossed with wild-type plants. All the F1 plants displayed a wild-type phenotype in osmotic stress response, and about three-quarters of the F2 progeny from self-pollinated F1 showed a wild-type phenotype (data not shown). The result suggests that ced1 is a recessive mutation in a single nuclear gene.

Wild-type and ced1 seedlings grown side-by-side on a half-strength Murashige and Skoog (MS) nutrient agar medium were measured for their ABA content under either normal or osmotic stress conditions. Under normal conditions, ABA in ced1 seedlings was found to be at a low level similar to that in the wild type (Figure 1A). To impose osmotic stress on the seedlings, 16-d-old seedlings grown in MS control plates were transferred to plates containing 40% PEG (average molecular weight 8000) solution. When incubated for 6 h in PEG solution, ABA levels in the wild-type seedlings increased dramatically. However, ABA levels in ced1 seedlings were considerably lower than in the wild type under the same treatment conditions (Figure 1A), indicating that the mutant was impaired in osmotic stress–induced ABA accumulation.

Figure 1.

Figure 1.

Open in a new tab

The ced1 Mutation Decreased ABA Accumulation and the Expression of ABA Biosynthesis and Signaling Genes in Response to Osmotic Stress Treatment.

(A) ABA contents in control or PEG-treated seedlings. Sixteen-day-old seedlings were transferred to plates containing 40% PEG solution for 6 h. Data are means ± se (n = 4 to 8). FW, fresh weight; WT, wild type.

(B) RNA gel blot analysis of a time course of NCED3 expression in ced1 and wild-type plants after transferring to 40% PEG. Twenty micrograms of total RNA were used per lane. rRNA was used as loading control.

(C) Transcript levels of ABA biosynthesis and signaling genes in response to osmotic stress in ced1 and wild-type seedlings. Sixteen-day-old plants grown on MS agar plates supplemented with 3% sucrose were transferred to plates containing PEG solutions of various concentrations and incubated for 1 h. Ethidium bromide–stained rRNAs were used as loading control.

In Arabidopsis, the NCED3 gene was induced by osmotic stress and the encoded enzyme is thought to control the level of ABA under osmotic stress (Iuchi et al., 2001). To ascertain whether the lower ABA accumulation in ced1 mutant plants under osmotic stress is a result of reduced NCED3 expression, we examined the steady state levels of the endogenous NCED3 transcript. Sixteen-day-old seedlings of wild-type and ced1 mutant plants were treated with 40% PEG solution for various time periods, and total RNA was extracted for RNA gel blot analysis. Indeed, the NCED3 transcript level was found to be substantially lower in the ced1 mutant than in the wild type (Figure 1B).

Osmotic Stress Induction of ABA Biosynthesis and Signaling Genes in ced1

Since ced1 was impaired in osmotic stress induction of NCED3 expression and ABA accumulation, we further determined the transcript levels of other ABA biosynthesis genes under osmotic stress conditions. These genes include ABA1 (encoding zeaxanthin epoxidase) (Marin et al., 1996) and AAO3 (encoding ABA aldehyde oxidase) (Seo et al., 2000) in addition to NCED3. Sixteen-day-old seedlings were treated with various concentrations of PEG for 1 h, and RNA gel blot analysis was performed. As shown in Figure 1C, the transcript levels of ABA1, NCED3, and AAO3 in both the wild type and ced1 were clearly upregulated by PEG-imposed osmotic stress, yet the levels decreased as the PEG concentrations further increased, perhaps as a result of severe damage to the seedlings. Nonetheless, the expression levels of these three genes were clearly lower in ced1 than in the wild type under the same treatment conditions. These data suggest that CED1 is required for full induction of multiple ABA biosynthesis genes by osmotic stress.

Since the ced1 mutation compromises stress induction of ABA biosynthesis genes, we were interested in testing whether it also affects the regulation of early ABA signaling genes. The ABA signaling pathway is mediated by ABA receptors in the PYR1/RCAR1 family, which, upon binding ABA, inhibit the type 2C protein phosphatases, such as ABI1 and ABI2 (Fujii et al., 2009; Ma et al., 2009; Park et al., 2009). ABI1 and ABI2 are negative regulators of the ABA signaling pathway (Gosti et al., 1999; Saez et al., 2004) and are induced by osmotic stress. Consistent with earlier observations, ABI1 and ABI2 were induced by osmotic stress in the wild type, and they were also induced in ced1 plants (Figure 1C). However, the expression levels of these two genes were substantially lower in ced1 than in the wild type, indicating that osmotic stress induction of these ABA signaling genes also requires normal CED1 functions (Figure 1C).

Reduced Expression of Osmotic Stress–Responsive Genes in ced1

We performed quantitative RT-PCR (qRT-PCR) to examine the expression levels of a number of osmotic stress–responsive genes, such as RD29A, KIN1, COR15A, RD22, RAB18, and P5CS1. These genes had low basal levels of expression in the absence of stress treatment but were strongly upregulated by PEG treatment in the wild type (40% for 6 h) (Figure 2A). In ced1 mutant plants, these genes showed no upregulation or much weaker upregulation in response to the same PEG treatment (Figure 2A). Interestingly, we also observed that expression of RD29A, RD22, COR15A, and COR47 was lower in the ced1 plants compared with the wild type in response to cold stress (see Supplemental Figure 2 online). These results suggest that CED1 may play an important role in regulating gene expression in response to osmotic stress and cold stress. To obtain a global view of the impact of ced1 mutation on gene regulation, we performed a microarray experiment using the Affymetrix ATH1 Genechips. RNA was extracted from both ced1 and wild-type seedlings that were treated with PEG (40%) for 6 h. Approximately 319 genes were found to have statistically significant changes in their expression levels in ced1 mutant compared with the wild type (see Supplemental Data Set 1 online). We categorized these genes into 19 functional groups using the gene ontology search tool (http://www.Arabidopsis.org/tools/bulk/go/index.jsp) with manual adjustment when necessary (see Supplemental Table 1 online). We observed an overrepresentation of genes related to ABA response (see Supplemental Figures 3 and 4 online). A majority of the genes that showed higher expression levels in ced1 under osmotic stress in our ATH1 array experiments (see Supplemental Data Set 1A online) are not responsive to ABA and are downregulated by osmotic stress in wild-type plants according to publicly available expression data (see Supplemental Figure 3 online). By contrast, most of the genes that showed lower expression levels in ced1 under osmotic stress in our ATH1 array experiments (see Supplemental Table 1B online) are upregulated by ABA or osmotic stress in wild-type plants according to publicly available expression data (see Supplemental Figure 4 online). Clearly, CED1 regulates ABA-responsive gene expression under osmotic stress.

Figure 2.

Figure 2.

Open in a new tab

Expression of Osmotic Stress–Responsive Genes in ced1 and Wild-Type Plants.

The qRT-PCR analysis was performed with total RNA from ced1 and wild-type (WT) plants under untreated control or osmotic stress treatment (40% PEG, 6 h) conditions. Real-time RT-PCR quantifications were normalized to the expression of TUB8. Error bars represent se from three independent experiments.

(A) qRT-PCR analysis of representative stress-responsive genes.

(B) Confirmation by qRT-PCR of the expression of genes showing significant changes between the wild type and the ced1 mutant in microarray analysis.

To validate the microarray results, we used qRT-PCR to analyze the expression of six genes that were found to show a significant expression difference in our microarray experiments. In agreement with the microarray data, the qRT-PCR analysis showed that At1g29395 (COR414-TM1), At3g05640 (PP2Cs), At5g15970 (KIN2), At5g52300 (RD29B), and At5g59320 (LTP3) were expressed at lower levels, while At5g45820 (PKS18) was expressed at a higher level in ced1 than in the wild type under the osmotic stress treatment (Figure 2B). There was no significant difference in the expression of these genes under the untreated control conditions. These results indicate that CED1 is important for osmotic stress induction of a large number of genes.

Germination and Seedling Growth of the ced1 Mutant Is More Sensitive to Osmotic Stress

The reduced osmotic stress induction of ABA biosynthesis genes and other osmotic stress-responsive genes in ced1 may affect stress tolerance of the mutant. We tested the sensitivity of seed germination and early seedling growth to osmotic stress. In the absence of osmotic stress, the germination of ced1 mutant seeds was similar to that of the wild type (Figure 3A). In the presence of osmotic stress (PEG-infused agar medium), seed germination of both ced1 and the wild type was delayed. However, the germination rates of ced1 seeds were substantially lower. For instance, at an osmotic potential of −0.7 MPa, only 19% of ced1 seeds, compared with 78% of the wild type, germinated. At −1.2 MPa, none of the mutant seeds could germinate, yet 21% of the wild-type seeds were still able to germinate (Figure 3A).

Figure 3.

Figure 3.

Open in a new tab

Osmotic Stress Phenotypes of the ced1 Mutant.

(A) Germination of wild-type and ced1 seeds in control or PEG-infused agar plates. Results are the average of four replicates ± se.

(B) Growth of wild-type (WT) and ced1 seedlings on control (left) or −0.5 MPa PEG-infused agar plates (right). Seeds were planted on shown plates, and the pictures were taken 4 weeks after seed imbibition.

(C) Wild-type and ced1 rosette plants were turgid immediately after detachment from roots, but ced1 plants became wilty 20 min later.

(D) Water loss rates of detached ced1 and wild-type shoots. The snrk2.6 mutant and its wild-type background ecotype Columbia (Col-0) are shown as controls. Data are means ± se (n = 4).

(E) ced1 plants were more sensitive to drought stress. Wild-type and ced1 plants were grown in soil with sufficient water for 3 weeks (Watered), and then water was withheld for 10 d (Drought) before rewatering. Plants were then allowed to recover for 2 d (Re-watered) before taking pictures. A representative picture for each treatment is shown.

(F) Quantification of the survival rate of the wild type and ced1 plants in (E). Survival rates and standard deviations were calculated from the results of four independent experiments.

In addition to inhibiting seed germination, osmotic stress also affects seedling growth. There is no obvious morphological difference between the wild type and ced1 when growing under control conditions in the agar plate (Figure 3B). At −0.5 MPa of osmotic potential in the medium, both shoot and root growth of ced1 seedlings was greatly inhibited, in contrast with the much milder impacts on wild-type seedlings (Figure 3B). These results indicate that ced1 mutant plants are more sensitive to osmotic stress during seed germination and early seedling development.

ced1 Plants Show Higher Transpiration Rates and Reduced Drought Resistance

When the aboveground parts were detached from roots at the rosette stage, the younger leaves of ced1 plants withered within 20 min under our room conditions (22 ± 2°C, ~30% RH). By contrast, wild-type leaves largely remained turgid under the same conditions (Figure 3C). This observation suggests that ced1 mutant plants may have a higher transpiration rate. Measurement of leaf water loss showed that ced1 mutant plants lost water much faster than wild-type plants (Figure 3D). Previous studies have shown that the snrk2.6/ost1 mutant was impaired in stomatal closure and had faster water loss rate than wild-type plants (Mustilli et al., 2002; Yoshida et al., 2002; Fujii et al., 2007). We found that the ced1 mutant had even higher water loss rate than snrk2.6 (Figure 3D). The greatly increased transpiration rate of ced1 leaves may affect drought resistance of the mutant.

To determine their drought resistance, wild-type and ced1 mutant plants were grown for 3 weeks in soil and then subjected to water withholding for an additional 10 d. At this time, ced1 mutant plants displayed obvious drought-stressed phenotypes, such as leaf wilting and senescence, whereas the wild-type plants were still turgid and their leaves remained green (Figure 3E). When the plants were rewatered and allowed to recover for 2 d, almost all ced1 mutant plants were dead, but the wild-type plants survived (Figure 3E). Quantitative analysis found that ~98 and 10% of wild-type and ced1 mutant plants survived the treatment, respectively (Figure 3F).

Map-Based Cloning of the CED1 Locus

The ced1 mutant phenotypes suggest a crucial role for CED1 in osmotic stress regulation of ABA biosynthesis and drought stress resistance. To determine the molecular nature of CED1, we took a map-based cloning approach to isolate the CED1 gene. A segregating F2 population was generated from a cross between ced1 (in the Columbia background) and the wild type Landsberg erecta. A total of 734 ced1 mutant plants were selected from the F2 population, and DNA was extracted from each plant for genetic mapping. CED1 was mapped to the lower arm of chromosome 1 between the simple sequence length polymorphism markers F23H11 and F5I14. Fine mapping within this chromosomal interval delimited the CED1 locus to an ~88-kb region within the BAC clone F1N19 (see Supplemental Figure 5A online). Within this region, 33 putative genes were predicted in The Arabidopsis Information Resource database (http://www.Arabidopsis.org/index.jsp). To find the mutation in ced1, 10 potential candidate open reading frames were amplified and sequenced. A single nucleotide mutation from G to A at position 1316 from the translation start site was found in ced1 mutant in At1g64670. This mutation would create a premature stop codon in the open reading frame that results in the truncation of the encoded protein (see Supplemental Figure 5B online).

The At1g64670 gene was annotated by the Arabidopsis Genome Initiative (http://www.Arabidopsis.org/index.jsp) as encoding an epidermis-expressed extracellular protein that likely functions as an α-β hydrolase. Interestingly, this locus was defined as BDG that controls epidermal cuticle development (Kurdyukov et al., 2006).

To confirm that the mutation in At1g64670 is responsible for the ced1 mutant phenotype, we transformed ced1 plants with an ~4.8-kb genomic fragment that includes the entire At1g64670 gene. Multiple transgenic lines were obtained, and two randomly chosen independent transformants were tested for their response to osmotic stress. Both transgenic lines were found to have osmotic stress tolerance similar to the wild type, whereas the original ced1 mutant seedlings were hypersensitive to the mild osmotic stress (−0.5 MPa) (Figure 4A). RNA gel blot analysis of these two lines confirmed the expression of the introduced gene (Figure 4B).

Figure 4.

Figure 4.

Open in a new tab

Functional Complementation of the ced1 Mutant and Expression of CED1/BDG in Response to Osmotic Stress or ABA.

(A) Complementation of ced1 by the wild-type CED1/BDG gene. The wild type (WT), ced1, and two homozygous ced1 transgenic lines (#1 and #3) expressing the wild type CED1. Plants were grown on MS (Control) or −0.5 MPa PEG-infused agar plates. The photograph was taken 4 weeks after seed imbibition.

(B) RNA gel blot analysis of the CED1/BDG gene expression level in the wild type, ced1, and the two homozygous transgenic lines (#1 and #3). rRNA was used as a loading control.

(C) The bdg-2 mutant was more sensitive to osmotic stress. Seeds were germinated and seedlings were grown on MS (Control) or −0.5 MPa PEG-infused agar plates for 4 weeks before taking the pictures.

(D) The ced1 mutant is allelic with bdg-2. Wild-type and F1 plants (ced1 × bdg-2) were grown on MS (Control) or −0.5 MPa PEG-infused agar plates for 4 weeks before taking the pictures.

(E) Regulation of CED1/BDG expression by osmotic stress and ABA. Sixteen-day-old plants were treated with 40% PEG or 100 μM ABA and incubated for the indicated time. 18S rRNA was used as a loading control.

We also examined the osmotic stress sensitivity of the bdg mutant. The bdg-2 allele contains a 7-bp deletion in the first exon (Kurdyukov et al., 2006) and is expected to be a null allele. When grown on agar plates with −0.5 MPa of water potential generated by PEG infusion, bdg-2 phenocopied the osmotic stress–sensitive phenotype of ced1 (Figure 4C). To provide further genetic proof that ced1 is allelic to bdg-2, we crossed bdg-2 with ced1. The resulting F1 seedlings were grown on −0.5 MPa PEG-infused agar plates. These seedlings exhibited increased osmotic stress sensitivity similar to the ced1 or bdg-2 single mutant, indicating a lack of complementation (Figure 4D; see Supplemental Table 2 online). Therefore, we renamed ced1 as bdg-5 (the 5th mutant allele of BDG). Collectively, these data indicate that At1g64670/BDG is CED1 and that CED1/BDG is required for osmotic stress tolerance.

Sensitivity of Other Cutin Biosynthesis Mutants to Osmotic Stress

The bdg mutants are characterized by irregular or interrupted cuticles, increased accumulation of cutin monomers and wax, as well as increased cuticular permeability (Kurdyukov et al., 2006). Although the enzymatic activity of BDG is unknown, the phenotypes of the bdg mutants suggest that BDG may be essential for the polymerization of cutin monomers at a late stage of cutin biosynthesis. While cuticles are well known to be able to minimize water loss and increase drought stress resistance, their role in osmotic stress tolerance is unclear. To ascertain whether the hypersensitivity to osmotic stress in bdg-5 is specific to the bdg-5 mutant or is a general consequence associated with the disruption of the cuticle, we sought to examine osmotic stress sensitivity in other cuticle mutants.

The cutin biosynthesis mutant aberrant induction of type three genes1 (att1-2) has a mutation in a cytochrome P450 monooxygenase that catalyzes fatty acid oxidation and has 30% of the wild-type cutin content (Xiao et al., 2004). The long-chain acyl-CoA synthetase (lacs2-1) mutant is impaired in cutin but not wax synthesis and is altered in the cuticle structure (Schnurr et al., 2004). The mutant may have a fivefold reduction in dicarboxylic acids, the typical monomers of Arabidopsis cutin (Bessire et al., 2007). LACERATA is also a cytochrome P450 monoxygenase (CYP86A8) important for the production of ω-hydroxy fatty acid components of the cutin polymer (Wellesen et al., 2001). The glycerol-3-phosphate acyltransferase double knockout mutant gpat4gpat8 has a 60 to 70% decrease in cutin monomer content, and the cuticle membrane in leaves is completely absent (Li et al., 2007). The relative positions of function of these enzymes in the cutin biosynthesis pathway are diagrammed in Figure 5A. Recently, overexpression of the putative transcription factor At MYB41 in Arabidopsis was found to abolish the expression of LACS2 and significantly reduced the expression of ATT1. Leaf permeability of the transgenic plants also significantly increased (Cominelli et al., 2008). A similar transgenic study also suggested that At MYB41 might negatively regulate short-term transcriptional responses to osmotic stress (Lippold et al., 2009). All of the above mutants have defects in cutin structures, with alterations either in the amount or composition of cutin. These mutants may or may not have changed wax contents. As controls, we also included two wax mutants, mah1-1 and cer7. The mah1-1 is mutated in the cytochrome P450 midchain alkane hydroxylase (CYP96A15) required for producing the wax components secondary alcohols and ketones (Greer et al., 2007). The cer7 mutation is in a putative 3′-5′ exoribonuclease of core exosome that degrades the transcript of a negative regulator of the wax gene CER3/WAX2/YRE (Hooker et al., 2007).

Figure 5.

Figure 5.

Open in a new tab

Osmotic Stress Sensitivity of Cutin Mutants.

(A) Schematic diagram of a possible cutin biosynthesis pathway. Shown are the synthesis of ω-hydroxy fatty acid–based cutin monomers and the enzymes that catalyze the reactions. The relative positions of these biosynthesis steps have not been completely determined (Pollard et al., 2008). CED1/BDG is proposed to function in the polymerization of the cutin monomers and is localized extracellularly. FFA, free fatty acid.

(B) Growth of wild-type (WT) and cutin mutant seedlings on MS (Control) or −0.5 MPa PEG-infused agar plates. Seeds were planted on the plates, and the pictures were taken 4 weeks after seed imbibition.

These mutants were grown under osmotic stress conditions, and their stress sensitivity was examined. In the absence of osmotic stress treatment, the postgermination growth of all mutants was similar to that of the wild type (Figure 5B). Under the PEG-imposed osmotic stress (−0.5 MPa), the growth of cutin mutants and the wild type were inhibited. However, the inhibition on all cutin mutants was much more severe than on the wild type (Figures 5B). The plants overexpressing At MYB41 (at-MYB41-OX) were also more sensitive to osmotic stress (see Supplemental Figure 6 online). These results indicated that these cutin-defective plants are much more sensitive to osmotic stress, similar to the bdg-5 mutant. By contrast, the sensitivity to osmotic stress of the wax mutants mah1-1 and cer7 was similar to that of the wild type (see Supplemental Figure 6 online).

Cutin but Not Wax Biosynthesis Mutants Are Impaired in Osmotic Stress Induction of ABA Biosynthesis and ABA Signaling Genes

The above data indicated that all of the cutin mutants examined are, like bdg-5, more sensitive to osmotic stress than the wild type. Since bdg-5 is defective in osmotic stress regulation of ABA biosynthesis genes as well as ABA signaling genes (Figures 1B and 1C), we asked whether the other cutin mutants are similarly impaired in osmotic stress regulation of these genes.

As shown in Figure 6, the transcript levels of ABA1, NCED3, ABI1, and ABI2 were clearly upregulated by osmotic stress in both the wild type and cutin mutants. Remarkably, the expression levels of these genes were all lower in the cutin mutants than in the wild type. Consistent with the negative roles of At MYB41 in regulating cutin biosynthesis, transgenic plants overexpressing At MYB41 showed a lower induction of ABA biosynthesis and signaling genes, similar to what was seen with other cutin biosynthesis mutants.

Figure 6.

Figure 6.

Open in a new tab

Steady State Transcript Levels of ABA Biosynthesis and Signaling Genes in Cutin or Wax Mutants.

Seedlings were treated with 40% PEG for the indicated time. Ten micrograms of total RNA were used per lane for the RNA gel blot analysis. rRNA was used as a loading control. Identities of probe sequences are indicated above the images, and the genotypes of different plants are indicated at left. Col-0, ecotype Columbia (wild-type plants).

The wax mutants mah1-1 and cer7 did not show any significant reduction in the expression of these ABA biosynthetic and signaling genes in response to osmotic stress (Figure 6). This is consistent with the unaltered sensitivity of the mutants to osmotic stress compared with the wild type (see Supplemental Figure 6 online). These data suggest that cutin but not wax biosynthesis mutants are unable to appropriately respond to osmotic stress in activating ABA biosynthesis and signaling genes, which may result in their increased sensitivity to even mild osmotic stress.

Increased Osmotic Sensitivity in bdg Mutant Plants Is Not Caused by Increased Permeability

Increased permeability was observed in aerial tissues of bdg-2 mutant plants (Kurdyukov et al., 2006; MacGregor et al., 2008). Similar to bdg-2, we found that aerial tissues of bdg-5 mutant plants show increased permeability (see Supplemental Figure 7 online). To examine whether the increased osmotic sensitivity in bdg-5 plants might be due to the increased permeability, we sowed the seeds in osmotic stress media immediately adjacent to strips of Parafilm as described (MacGregor et al., 2008). Under this condition, the aerial tissues will grow only onto the Parafilm and will not be in contact with the medium. Consistent with the previous observation in bdg-2 (MacGregor et al., 2008), lateral root formation in bdg-5 is reduced with the use of Parafilm in control media (see Supplemental Figure 8A online). However, the bdg-5 seedlings remain hypersensitive to mild osmotic stress imposed by PEG-infused agar media (see Supplemental Figure 8A online). These results suggest that increased osmotic stress sensitivity in bdg-5 plants is not caused by increased aerial tissue permeability. We observed similar results in bdg-2 mutant when seeds of the bdg-2 were germinated next to strips of Parafilm in the presence of osmotic stress (see Supplemental Figure 8B online). We also tested another mutant, gpat5, which is defective in glycerol-3-phosphate acyltransferase. The gpat5 mutant displayed severalfold reduction in suberin in its seed coats and but no change in surface wax (Beisson et al., 2007). Seed coats of gpat5 mutant plants had a great increase in permeability (Beisson et al., 2007). We found that seed germination and postgermination seedling development of gpat5 are not affected by osmotic stress conferred by PEG (see Supplemental Figure 8C online). The result suggests that increased seed coat permeability does not necessarily lead to increased osmotic stress sensitivity in germination. Together, the above data support that increased permeability is not the cause of hypersensitivity of bdg mutants to osmotic stress conferred by PEG.

BDG Expression Is Induced by Osmotic Stress and ABA

The critical role of BDG in osmotic stress tolerance and ABA biosynthesis prompted us to investigate how the BDG gene is regulated in response to osmotic stress and ABA. RNA gel blot analysis was performed with total RNA obtained from the wild-type seedlings grown on the MS agar media for 16 d. As shown in Figure 4E, the transcript level of BDG was substantially upregulated after 1 h of osmotic stress but returned to the untreated control level after prolonged stress. ABA induced a higher expression level of BDG, which peaked at 3 h after the treatment and gradually decreased thereafter.

DISCUSSION

Plant aerial parts are covered with a layer of hydrophobic cuticle that prevents the epidermal cells from direct contact with the dry environment of the air. This physical barrier minimizes water loss, discourages pathogens and herbivores, and protects the cells below from damage by UV and other environmental stresses (Kolattukudy, 2001; Kosma and Jenks, 2007; Pollard et al., 2008). The cuticle also prevents organ fusion (Sieber et al., 2000) and is thus required for normal morphogenesis of the aerial organs. Although the cuticle has been known to play important roles as a physical barrier to enhance plant resistance to abiotic stress and cutin was recently shown to be important for plant resistance to pathogens, a role for cuticle in regulating ABA biosynthesis and osmotic stress resistance was unexpected.

Cutin and Cuticle Are Responsible for the Defects in Osmotic Responses in bdg-5 and Other Cuticle Mutants

With the critical role of CED1 in regulating NCED3 gene expression, one may expect CED1 to encode a typical regulatory component such as a transcriptional factor. Unexpectedly, CED1 was found to be allelic to BGD, which encodes a putative α/β hydrolase fold protein recently shown to be essential for cutin synthesis. Although the catalytic activity of this putative enzyme is unknown, its tissue-specific expression and localization (epidermis and extracellular) and the mutant phenotypes (accumulation of cutin monomers and waxes) support that CED1/BDG may be an enzyme that catalyzes either the polymerization of cutin monomers or some yet unknown process in the cutin biosynthesis pathway (Kurdyukov et al., 2006), although the possibility that BDG could directly be a signaling component cannot be completely ruled out.

Unlike BDG, whose catalytic activity is still unknown, most of the other cutin biosynthesis loci encode enzymes belonging to well-defined catalytic groups, although their detailed reactions and their relative positions in the overall cutin biosynthesis pathway may not be clear. For instance, ATT1 is a cytochrome P450 monooxygenase (CYP86A2) catalyzing ω-oxidation of fatty acids and the att1 mutant has 30% of the wild-type cutin level (Xiao et al., 2004). LACS2 is a long-chain acyl-CoA synthase that produces ω-hydroxy fatty acyl-CoA for cutin synthesis, and its mutation reduces the thickness of the cutin membrane (Schnurr et al., 2004). Our finding that all these cutin biosynthesis mutants are similarly impaired in osmotic stress induction of ABA biosynthethic genes and osmotic stress tolerance indicates that most likely it is the cutin integrity that is responsible for the defects in osmotic stress responses. Since these mutants are affected in different steps of cutin biosynthesis, it is less likely that a common substrate or product in cutin biosynthesis acts as a signal for the activation of ABA biosynthesis.

While it has long been accepted that plant cuticles function in increasing drought resistance by minimizing nonstomatal transpiration water loss, because the cuticle constitutes a physical water-repelling barrier to prevent water escape, it is unexpected that the cuticle may play an active role in regulating ABA biosynthesis and osmotic stress resistance. The exact mechanism for cuticle regulation of osmotic stress activation of ABA biosynthesis and other stress responses is unknown at this time. However, our study clearly indicates that an intact cuticle is essential for osmotic stress to activate ABA biosynthesis as well as for plants to develop osmotic stress resistance, providing an unexpected link between cuticle integrity and cell signaling. This discovery may have some resemblance to the cell wall integrity response in yeast, where cell wall damages elicit a signaling cascade that activates cell wall remodeling-related genes (Levin, 2005). Recently, it was also found that the high osmolarity glycerol pathway is also involved in cell wall damage response (Bermejo et al., 2008; Garcia et al., 2009), which implicated a connection between cell wall integrity and osmotic stress response in yeast cells.

Possible Connections between Cuticle and Cell Wall in Osmotic Stress Regulation of ABA Biosynthesis

It is well recognized that the growth of a plant cell is largely determined by the balance between turgor pressure from the protoplasts and the constriction imposed by the cell wall. The direction for signaling cell growth is from within the cell to the cell wall so that the wall can be loosened and expanded. Whether and how the cell wall may send a retrograde signal to the cell to make appropriate adjustment to cope with external osmotic stress is currently unknown. Although the conventional idea is that cellular responses to osmotic stress start with the perception of the stress at the plasma membrane, recently, however, several genetic studies revealed connections between the cell wall and osmotic stress responses. Mutations in a cellulose synthase-like gene SOS6/CSLD5 caused the mutants to be hypersensitive to osmotic stress, although the cell wall composition of this mutant was only subtly affected (Zhu et al., 2010). A mutation in the cellulose synthase gene CesA8/IRX1, on the other hand, led to increased osmotic stress tolerance in the mutant (Chen et al., 2005). These studies suggest a critical role for cell wall in osmotic stress tolerance, although the mechanisms involved are unknown.

A role for the cell wall in osmotic stress induction of ABA biosynthesis has also been implicated in several studies. It seems that a net change (reduction) in turgor pressure is required for osmotic stress induction of ABA biosynthesis in detached leaves (Pierce and Raschke, 1980). Penetrating solutes equally lower the osmotic potential of the cell but failed to elicit ABA biosynthesis (Creelman and Zeevaart, 1985). Similarly, osmotic stress treatments failed to increase ABA biosynthesis in Vicia faba (Lahr and Raschke, 1988) or barley (Hordeum vulgare) protoplasts (Loveys and Robinson, 1987), but the same treatments could activate ABA biosynthesis in leaf discs in these studies. Nonetheless, one other study reported that ABA biosynthesis could occur in protoplasts of rose petals or Amaranthus leaves (Bianco-Trinchant and Le Page-Degivry, 1998). Aside from the difference in plant materials and experimental conditions, it is not clear why these research results differ.

Unlike a complete lack or a general compromise of the cell wall integrity as seen with protoplasts or the cell wall mutants in previous studies, ced1 mutant seedlings may not have a global defect in the cell wall structure. Gene expression and immunolocalization assays indicated that BDG is expressed exclusively in epidermis and the protein is localized extracellularly (Kurdyukov et al., 2006). Consistent with this pattern of localization, the structural defects of bdg-5 appear to be very specific and restricted to the cuticles and epidermis, although the mutant does have other developmental defects. The isolation of the ced1 mutant thus defines a specialized and unexpected structure (i.e., the cuticle) as essential for osmotic stress sensing and response.

Cutin but Not Wax Is Required for Osmotic Stress Response and Tolerance

The plant cuticle is mainly composed of cutin embedded with intracuticular waxes and covered with epicuticular waxes. Cutin is a polyester of hydroxy and epoxy-hydroxy C16 or C18 fatty acids and glycerol, while waxes consist of very-long-chain fatty acids with chain length from 24 to 26 carbons along with various derivations and modifications. Although both components work together structurally and functionally to seal the epidermis and prevent water loss (Kosma and Jenks, 2007), our studies revealed that they have very different roles in osmotic stress responses. Unlike bdg-5 and other cutin mutants, mutants defective in wax biosynthesis are not affected in either the expression of ABA biosynthesis and signaling genes (Figure 6) or osmotic stress resistance (see Supplemental Figure 6 online). These results suggest that the defect in cutin but not wax biosynthesis affects cuticle’s ability to respond to osmotic stress. In fact, in several cutin biosynthesis mutants, including bdg-5, wax contents actually increased (Schnurr et al., 2004; Kurdyukov et al., 2006; Bessire et al., 2007), perhaps as a result of increased channeling of early common substrates to the wax biosynthesis pathway or the activation of wax biosynthetic genes to compensate for the loss of cutin in cuticles.

Does Osmotic Stress Regulation of ABA Biosynthesis Contribute to the Altered Disease Resistance in Cutin Biosynthesis Mutants?

The discovery that cutin and cuticles are required for either sensing or responding to osmotic stress is very intriguing. Recently, a role of cutin biosynthesis and cuticle integrity in disease resistance has emerged. In multiple mutant screens for resistance to bacterial or fungal pathogens, several mutants susceptible to bacterial pathogens were found to have defects in cutin biosynthesis or integrity (Xiao et al., 2004; Tang et al., 2007). Interestingly, these mutants are more resistant to the necrotrophic fungus Botrytis cinerea (Bessire et al., 2007; Chassot et al., 2007; Tang et al., 2007), although the mechanisms for this immunity are unknown. Cutin monomers or other constituents of the cuticles could elicit reactive oxygen species production (Fauth et al., 1998) and can potentially serve as signals to activate defense responses in the host. It is known that ABA is actively involved in plant resistance to pathogens (Mauch-Mani and Mauch, 2005; Adie et al., 2007; de Torres-Zabala et al., 2007; Fan et al., 2009); whether reduced ABA biosynthesis in cuticle mutants has a direct role in regulating disease resistance will need to be investigated in the future.

Possible Mechanisms for Cuticle Regulation of ABA Biosynthesis

At present, we can only speculate on why cuticle integrity would be needed for osmotic stress induction of ABA biosynthesis and osmotic stress tolerance. Aside from possible signaling molecules derived from cutin biosynthesis, there may be other epidermal cell wall– or even cuticle-associated proteins that are essential for the sensing of osmotic stress signals. Many wall-associated putative receptors, sensors, and other proteins exist, yet their functions in cell signaling are still largely unknown (Humphrey et al., 2007). An intact cuticle may be needed for the correct localization or function of these signaling molecules. BDG or other cutin biosynthesis enzymes themselves may not be such molecules since these diverse biosynthetic enzymes have little structural or functional similarity to signaling molecules that might directly serve a signaling role in osmotic stress response.

Alternatively, the physical integrity of the cuticle may be required for osmotic stress responses. Under normal growth conditions, plant cells are under strong positive pressure from the cell wall (turgor pressure). Under osmotic stress, turgor pressure is reduced due to decreased water potential in the apoplast, and the cell may sense this change in turgor pressure and trigger a signal transduction cascade to activate ABA biosynthesis and other osmotic stress responses. When the cuticle is disrupted and the epidermis cell wall may become less rigid and more flexible, the mutant cell wall may no longer be able to sense changes in the osmotic potential of the cell. This would result in no response or reduced response to osmotic stress as seen in ced1 and other cutin biosynthesis mutants.

One intriguing fact about BDG is that the gene and protein are mainly expressed in the epidermis (Kurdyukov et al., 2006), while ABA biosynthesis enzymes are found mainly associated with the vascular tissues (Cheng et al., 2002; Gonzalez-Guzman et al., 2002; Tan et al., 2003; Koiwai et al., 2004). It is yet unclear how the signal perceived at the epidermal cells would be transmitted to the vascular tissues. Likely cuticle integrity may impact not only the epidermal cells underneath but also the leaf and other organs of the plant as a whole. In addition to leaf epidermal cells, BDG as a putative enzyme in the cutin biosynthesis pathway was expressed in root epidermis and probably pericycle cells where lateral roots emerge (Kurdyukov et al., 2006). Cutin is biochemically similar to the suberin found in root endodermis of most higher plants, and some of their biosynthesis pathways may overlap (Franke et al., 2005), although it is not clear whether bdg-5 has an altered suberin. Thus, one additional possible connection between cuticle biosynthesis and osmotic stress–induced ABA biosynthesis may have to do with the coupling of suberin integrity and ABA biosynthesis in root vascular tissues. These possibilities would need to be tested in future experiments.

METHODS

Plant Materials and Growth Conditions

A firefly LUC reporter gene driven by the drought stress-responsive NCED3 promoter (−1083 to + 76 bp from the transcription start site) was introduced into Arabidopsis thaliana plants in the Columbia glabrous1 background. Seeds from one homozygous line expressing a single functional copy of the NCED3:LUC gene (referred to as the wild type) were mutagenized with ethyl methanesulfonate. The ced1 mutant with altered NCED3:LUC gene expression was isolated from M2 seedlings with a CCD camera imaging system (Ishitani et al., 1997).

The att1-1 and att1-2 mutants were kindly provided by Jian-Min Zhou (National Institute of Biological Sciences, Beijing). The lacs2-1 mutant was kindly provided by John Browse (Washington State University). lcr and bdg were kindly provided by Alexander Yephremov (Max Planck Institute for Plant Breeding Research, Germany). The gpat4 gpat8 double mutant and gpat5 were kindly provided by John Ohlrogge (Michigan State University). The At MYB41 overexpression line was kindly provided by Chiara Tonelli (Università degli Studi di Milano, Italy). The mah1-1 (SALK_049943), cer7 (CS8017), and snrk2.6 (SALK_008068) T-DNA insertion lines were obtained from the ABRC (Columbus, OH). Arabidopsis seedlings on MS medium agar plates (1× MS salts, 3% Suc, and 1.5% agar, pH 5.7) were routinely grown under continuous white light (~75 μmol m−2 s−1) at 23 ± 1°C. Soil-grown plants were kept under a 16-h-light/8-h-dark photoperiod at 23 ± 1°C.

Physiological Assays

For germination assays, seeds were planted on either a half-strength MS 1.5% agar medium with 3% Suc buffered with 2 mM MES or PEG-infused agar plates without Suc. In each experiment, at least 100 seeds per genotype were stratified at 4°C for 3 d, and radicle emergence was scored as being germinated at the indicated time.

For leaf water loss measurements, fully expanded leaves were removed from 4-week-old plants and incubated under the same conditions used for seedling growth, and each sample (consisting of three to four individual leaves) was weighed at the indicated time. For drought treatment, plants were grown in soil with sufficient water for 3 weeks, and then the water was withheld for durations as indicated.

For ABA quantification, ABA was measured by radioimmunoassay as previously described (Verslues et al., 2007).

Genetic Mapping and Complementation

The ced1 mutant was crossed with the Landsberg erecta accession, and 734 mutant plants were chosen from the F2 generation by their osmotic stress sensitivity phenotype. Simple sequence length polymorphism markers were designed according to the information in the Cereon Arabidopsis Polymorphism Collection and used to analyze recombination events (Jander et al., 2002). The ced1 mutation was first mapped to chromosome 1 between F23H11 and F5I14. Fine mapping within this chromosomal interval narrowed the CED1 locus to an ~88-kb region within the BAC clone F1N19. All candidate genes in this region were sequenced from the ced1 mutant and compared with those in GenBank to find the ced1 mutation.

For complementation of the ced1 mutant, an ~4.8-kb genomic fragment that includes 1813 bp upstream of the translation initiation codon and 549 bp downstream of the translation stop codon was amplified (see Supplemental Table 3 online for primer sequences). The amplified fragment was cloned into the pCAMBIA 1380 vector. The construct was transferred into Agrobacterium tumefaciens (GV3101 strain), and plants were transformed using the floral dip method (Clough and Bent, 1998).

RNA Gel Analysis and Real-Time RT-PCR Analysis

Sixteen-day-old seedlings grown on MS media (1× MS salts, 3% Suc, and 0.6% agar, pH 5.7) plates that were placed in a horizontal orientation were used for RNA gel analysis. Total RNA was extracted from the wild type, mutants, and transgenic plants with the Trizol reagent (Invitrogen). Total RNA was separated on a 1.2% formaldehyde-MOPS agarose gel. The blots were probed, washed, and wrapped in plastic wrap to keep them from drying out and then immediately exposed to x-ray film for autoradiography as described (Xiong et al., 2001).

For real-time RT-PCR analysis, 5 μg of total RNA isolated with the RNeasy plant mini kit (Qiagen) was used for the first-strand cDNA synthesized using SuperScript III first-strand synthesis supermix (Invitrogen). The cDNA reaction mixture was diluted three times, and 5 μL was used as a template in a 25-μL PCR reaction. PCR was performed after a preincubation at 95°C for 3 min and followed by 40 cycles of denaturation at 95°C for 15 s, annealing at 55°C for 40 s, and extension at 72°C for 40 s. All the reactions were performed in the iQ5 real-time PCR detection system using iQ SYBR Green Supermix (Bio-Rad). Each experiment was replicated three times. The comparative Ct method was applied. The primers used in this study are listed in Supplemental Table 3 online.

Microarray Analysis

For Affymetrix GeneChip array analysis, wild-type and ced1 seedlings were grown on MS plates for 15 d at 22°C with 16 h light and 8 h darkness. Total RNA was extracted using an RNeasy plant mini kit (Qiagen) and used for preparation of biotin-labeled complementary RNA targets. Microarray analysis was performed as described by Breitling et al. (2004). Two biological replicates were used for each genotype. We applied the Robust Multiarray Averaging normalization method for our data sets. The RMA method for computing an expression measure begins by computing background-corrected perfect match intensities for each perfect match cell on every GeneChip. The normalized data were further analyzed, and P values were generated by affymGUI component of Bioconductor in statistics environment R, with the default parameters (Irizarry et al., 2003; Gentleman et al., 2004). We selected lists of genes with statistically significant changes between mutant and wild type under osmotic stress conditions by the RankProd method (nonparamteric method for identifying differentially expressed [up- or downregulated] genes based on the estimated percentage of false predictions [pfp]) (Hong et al., 2006) (pfp < 0.05). RankProd results were summarized with the script written in PERL.

Accession Numbers

Sequence data from this article can be found in the Arabidopsis Genome Initiative or GenBank/EMBL databases under the following accession numbers: BDG/CED1 (At1g64670), NCED3 (At3g14440), ABA1 (At5g67030), AAO3 (At2g27150), ABI1 (At4g26080), ABI2 (At5g57050), RD29A (At5g52310), KIN1 (At5g15960), COR15A (At2g42540), RD22 (At5g25610), RAB18 (At5g66400), P5CS1 (At2g39800), COR414-TM1 (At1g29395), PP2Cs (At3g05640), KIN2 (At5g15970), RD29B (At5g52300), LTP3 (At5g59320), and PKS18 (At5g45820). The microarray data discussed in this publication have been deposited in the National Center for Biotechnology Information’s Gene Expression Omnibus (Edgar et al., 2002) and are accessible through GEO Series accession number GSE 25889 (http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE 25889).

Supplemental Data

The following materials are available in the online version of this article.

Acknowledgments

We thank Paul E. Verslues for help with the initial phase of the work, Iida Kei for help with analyzing the microarray data, and Rebecca Stevenson for technical assistance. This work was supported by National Institutes of Health Grants R01GM070795 and R01GM059138 to J.-K.Z. and by National Science Foundation Grant IOS0919745 to J.Z.

References

  1. Adie B.A., Perez-Perez J., Perez-Perez M.M., Godoy M., Sanchez-Serrano J.J., Schmelz E.A., Solano R. (2007). ABA is an essential signal for plant resistance to pathogens affecting JA biosynthesis and the activation of defenses in Arabidopsis. Plant Cell 19: 1665–1681 [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Beisson F., Li Y., Bonaventure G., Pollard M., Ohlrogge J.B. (2007). The acyltransferase GPAT5 is required for the synthesis of suberin in seed coat and root of Arabidopsis. Plant Cell 19: 351–368 [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Bermejo C., Rodriguez E., Garcia R., Rodriguez-Pena J.M., Rodriguez de la Concepcion M.L., Rivas C., Arias P., Nombela C., Posas F., Arroyo J. (2008). The sequential activation of the yeast HOG and SLT2 pathways is required for cell survival to cell wall stress. Mol. Biol. Cell 19: 1113–1124 [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Bessire M., Chassot C., Jacquat A.C., Humphry M., Borel S., Petetot J.M., Metraux J.P., Nawrath C. (2007). A permeable cuticle in Arabidopsis leads to a strong resistance to Botrytis cinerea. EMBO J. 26: 2158–2168 [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bianco-Trinchant J., Le Page-Degivry M.T. (1998). ABA synthesis in protoplasts of different origin in response to osmotic stress. Plant Growth Regul. 25: 135–141 [Google Scholar]
  6. Borsani O., Cuartero J., Valpuesta V., Botella M.A. (2002). Tomato tos1 mutation identifies a gene essential for osmotic tolerance and abscisic acid sensitivity. Plant J. 32: 905–914 [DOI] [PubMed] [Google Scholar]
  7. Breitling R., Armengaud P., Amtmann A., Herzyk P. (2004). Rank products: A simple, yet powerful, new method to detect differentially regulated genes in replicated microarray experiments. FEBS Lett. 573: 83–92 [DOI] [PubMed] [Google Scholar]
  8. Chassot C., Nawrath C., Metraux J.P. (2007). Cuticular defects lead to full immunity to a major plant pathogen. Plant J. 49: 972–980 [DOI] [PubMed] [Google Scholar]
  9. Chen Z., Hong X., Zhang H., Wang Y., Li X., Zhu J.K., Gong Z. (2005). Disruption of the cellulose synthase gene, AtCesA8/IRX1, enhances drought and osmotic stress tolerance in Arabidopsis. Plant J. 43: 273–283 [DOI] [PubMed] [Google Scholar]
  10. Cheng W.H., Endo A., Zhou L., Penney J., Chen H.C., Arroyo A., Leon P., Nambara E., Asami T., Seo M., Koshiba T., Sheen J. (2002). A unique short-chain dehydrogenase/reductase in Arabidopsis glucose signaling and abscisic acid biosynthesis and functions. Plant Cell 14: 2723–2743 [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Clough S.J., Bent A.F. (1998). Floral dip: A simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J. 16: 735–743 [DOI] [PubMed] [Google Scholar]
  12. Cominelli E., Sala T., Calvi D., Gusmaroli G., Tonelli C. (2008). Over-expression of the Arabidopsis AtMYB41 gene alters cell expansion and leaf surface permeability. Plant J. 53: 53–64 [DOI] [PubMed] [Google Scholar]
  13. Creelman R.A., Zeevaart J.A. (1985). Abscisic acid accumulation in spinach leaf slices in the presence of penetrating and nonpenetrating solutes. Plant Physiol. 77: 25–28 [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Cutler S.R., Rodriguez P.L., Finkelstein R.R., Abrams S.R. (2010). Abscisic acid: Emergence of a core signaling network. Annu. Rev. Plant Biol. 61: 651–679 [DOI] [PubMed] [Google Scholar]
  15. de Torres-Zabala M., Truman W., Bennett M.H., Lafforgue G., Mansfield J.W., Rodriguez Egea P., Bogre L., Grant M. (2007). Pseudomonas syringae pv. tomato hijacks the Arabidopsis abscisic acid signalling pathway to cause disease. EMBO J. 26: 1434–1443 [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Edgar R., Domrachev M., Lash A.E. (2002). Gene Expression Omnibus: NCBI gene expression and hybridization array data repository. Nucleic Acids Res. 30: 207–210 [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Fan J., Hill L., Crooks C., Doerner P., Lamb C. (2009). Abscisic acid has a key role in modulating diverse plant-pathogen interactions. Plant Physiol. 150: 1750–1761 [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Fauth M., Schweizer P., Buchala A., Markstadter C., Riederer M., Kato T., Kauss H. (1998). Cutin monomers and surface wax constituents elicit H2O2 in conditioned cucumber hypocotyl segments and enhance the activity of other H2O2 elicitors. Plant Physiol. 117: 1373–1380 [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Finkelstein R.R., Gampala S.S., Rock C.D. (2002). Abscisic acid signaling in seeds and seedlings. Plant Cell 14(suppl.): S15–S45 [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Finkelstein R.R., Rock C.D. (2008). Abscisic acid biosynthesis and response. In The Arabidopsis Book 1: e0058, doi/10.1199/tab.0058 [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Franke R., Briesen I., Wojciechowski T., Faust A., Yephremov A., Nawrath C., Schreiber L. (2005). Apoplastic polyesters in Arabidopsis surface tissues–A typical suberin and a particular cutin. Phytochemistry 66: 2643–2658 [DOI] [PubMed] [Google Scholar]
  22. Fujii H., Chinnusamy V., Rodrigues A., Rubio S., Antoni R., Park S.Y., Cutler S.R., Sheen J., Rodriguez P.L., Zhu J.K. (2009). In vitro reconstitution of an abscisic acid signalling pathway. Nature 462: 660–664 [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Fujii H., Verslues P.E., Zhu J.K. (2007). Identification of two protein kinases required for abscisic acid regulation of seed germination, root growth, and gene expression in Arabidopsis. Plant Cell 19: 485–494 [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Fujii H., Zhu J.K. (2009). Arabidopsis mutant deficient in 3 abscisic acid-activated protein kinases reveals critical roles in growth, reproduction, and stress. Proc. Natl. Acad. Sci. USA 106: 8380–8385 [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Garcia R., Rodriguez-Pena J.M., Bermejo C., Nombela C., Arroyo J. (2009). The high osmotic response and cell wall integrity pathways cooperate to regulate transcriptional responses to zymolyase-induced cell wall stress in Saccharomyces cerevisiae. J. Biol. Chem. 284: 10901–10911 [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Gentleman R.C., et al. (2004). Bioconductor: open software development for computational biology and bioinformatics. Genome Biol. 5: R80. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Gonzalez-Guzman M., Apostolova N., Belles J.M., Barrero J.M., Piqueras P., Ponce M.R., Micol J.L., Serrano R., Rodriguez P.L. (2002). The short-chain alcohol dehydrogenase ABA2 catalyzes the conversion of xanthoxin to abscisic aldehyde. Plant Cell 14: 1833–1846 [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Gosti F., Beaudoin N., Serizet C., Webb A.A., Vartanian N., Giraudat J. (1999). ABI1 protein phosphatase 2C is a negative regulator of abscisic acid signaling. Plant Cell 11: 1897–1910 [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Greer S., Wen M., Bird D., Wu X., Samuels L., Kunst L., Jetter R. (2007). The cytochrome P450 enzyme CYP96A15 is the midchain alkane hydroxylase responsible for formation of secondary alcohols and ketones in stem cuticular wax of Arabidopsis. Plant Physiol. 145: 653–667 [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Hong F., Breitling R., McEntee C.W., Wittner B.S., Nemhauser J.L., Chory J. (2006). RankProd: A bioconductor package for detecting differentially expressed genes in meta-analysis. Bioinformatics 22: 2825–2827 [DOI] [PubMed] [Google Scholar]
  31. Hooker T.S., Lam P., Zheng H., Kunst L. (2007). A core subunit of the RNA-processing/degrading exosome specifically influences cuticular wax biosynthesis in Arabidopsis. Plant Cell 19: 904–913 [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Humphrey T.V., Bonetta D.T., Goring D.R. (2007). Sentinels at the wall: Cell wall receptors and sensors. New Phytol. 176: 7–21 [DOI] [PubMed] [Google Scholar]
  33. Irizarry R.A., Hobbs B., Collin F., Beazer-Barclay Y.D., Antonellis K.J., Scherf U., Speed T.P. (2003). Exploration, normalization, and summaries of high density oligonucleotide array probe level data. Biostatistics 4: 249–264 [DOI] [PubMed] [Google Scholar]
  34. Ishitani M., Xiong L., Stevenson B., Zhu J.-K. (1997). Genetic analysis of osmotic and cold stress signal transduction in Arabidopsis thaliana: Interactions and convergence of abscisic acid-dependent and abscisic acid-independent pathways. Plant Cell 9: 1935–1949 [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Iuchi S., Kobayashi M., Taji T., Naramoto M., Seki M., Kato T., Tabata S., Kakubari Y., Yamaguchi-Shinozaki K., Shinozaki K. (2001). Regulation of drought tolerance by gene manipulation of 9-cis-epoxycarotenoid dioxygenase, a key enzyme in abscisic acid biosynthesis in Arabidopsis. Plant J. 27: 325–333 [DOI] [PubMed] [Google Scholar]
  36. Jander G., Norris S.R., Rounsley S.D., Bush D.F., Levin I.M., Last R.L. (2002). Arabidopsis map-based cloning in the post-genome era. Plant Physiol. 129: 440–450 [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Koiwai H., Nakaminami K., Seo M., Mitsuhashi W., Toyomasu T., Koshiba T. (2004). Tissue-specific localization of an abscisic acid biosynthetic enzyme, AAO3, in Arabidopsis. Plant Physiol. 134: 1697–1707 [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Kolattukudy P.E. (2001). Polyesters in higher plants. Adv. Biochem. Eng. Biotechnol. 71: 1–49 [DOI] [PubMed] [Google Scholar]
  39. Kosma D.K., Jenks M.A. (2007). Eco-physiological and molecular-genetic determinants of plant cuticle function in drought and salt stress tolerance. In Advances in Molecular Breeding toward Drought and Salt Tolerant Crops. Jenks M.A., Hasegawa P.M., Jain S.M., eds (Dordrecht, The Netherlands: Springer; ), pp. 91–120 [Google Scholar]
  40. Kurdyukov S., Faust A., Nawrath C., Bar S., Voisin D., Efremova N., Franke R., Schreiber L., Saedler H., Metraux J.P., Yephremov A. (2006). The epidermis-specific extracellular BODYGUARD controls cuticle development and morphogenesis in Arabidopsis. Plant Cell 18: 321–339 [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Lahr W., Raschke K. (1988). Abscisic acid contents and concentrations in protoplasts from guard cells and mesophyll cells of Vicia faba L. Planta 173: 528–531 [DOI] [PubMed] [Google Scholar]
  42. Levin D.E. (2005). Cell wall integrity signaling in Saccharomyces cerevisiae. Microbiol. Mol. Biol. Rev. 69: 262–291 [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Li Y., Beisson F., Koo A.J., Molina I., Pollard M., Ohlrogge J. (2007). Identification of acyltransferases required for cutin biosynthesis and production of cutin with suberin-like monomers. Proc. Natl. Acad. Sci. USA 104: 18339–18344 [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Lippold F., Sanchez D.H., Musialak M., Schlereth A., Scheible W.R., Hincha D.K., Udvardi M.K. (2009). AtMyb41 regulates transcriptional and metabolic responses to osmotic stress in Arabidopsis. Plant Physiol. 149: 1761–1772 [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Loveys B.R., Robinson S.P. (1987). Abscisic acid synthesis and metabolism in barley leaves and protoplasts. Plant Sci. 49: 23–30 [Google Scholar]
  46. Ma Y., Szostkiewicz I., Korte A., Moes D., Yang Y., Christmann A., Grill E. (2009). Regulators of PP2C phosphatase activity function as abscisic acid sensors. Science 324: 1064–1068 [DOI] [PubMed] [Google Scholar]
  47. MacGregor D.R., Deak K.I., Ingram P.A., Malamy J.E. (2008). Root system architecture in Arabidopsis grown in culture is regulated by sucrose uptake in the aerial tissues. Plant Cell 20: 2643–2660 [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Marin E., Nussaume L., Quesada A., Gonneau M., Sotta B., Hugueney P., Frey A., Marion-Poll A. (1996). Molecular identification of zeaxanthin epoxidase of Nicotiana plumbaginifolia, a gene involved in abscisic acid biosynthesis and corresponding to the ABA locus of Arabidopsis thaliana. EMBO J. 15: 2331–2342 [PMC free article] [PubMed] [Google Scholar]
  49. Mauch-Mani B., Mauch F. (2005). The role of abscisic acid in plant-pathogen interactions. Curr. Opin. Plant Biol. 8: 409–414 [DOI] [PubMed] [Google Scholar]
  50. Mustilli A.C., Merlot S., Vavasseur A., Fenzi F., Giraudat J. (2002). Arabidopsis OST1 protein kinase mediates the regulation of stomatal aperture by abscisic acid and acts upstream of reactive oxygen species production. Plant Cell 14: 3089–3099 [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Nambara E., Marion-Poll A. (2005). Abscisic acid biosynthesis and catabolism. Annu. Rev. Plant Biol. 56: 165–185 [DOI] [PubMed] [Google Scholar]
  52. Okamoto M., Tanaka Y., Abrams S.R., Kamiya Y., Seki M., Nambara E. (2009). High humidity induces abscisic acid 8'-hydroxylase in stomata and vasculature to regulate local and systemic abscisic acid responses in Arabidopsis. Plant Physiol. 149: 825–834 [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Park S.Y., et al. (2009). Abscisic acid inhibits type 2C protein phosphatases via the PYR/PYL family of START proteins. Science 324: 1068–1071 [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Pierce M., Raschke K. (1980). Correlation between loss of turgor and accumulation of abscisic acid in detached leaves. Planta 148: 174–182 [DOI] [PubMed] [Google Scholar]
  55. Pollard M., Beisson F., Li Y., Ohlrogge J.B. (2008). Building lipid barriers: Biosynthesis of cutin and suberin. Trends Plant Sci. 13: 236–246 [DOI] [PubMed] [Google Scholar]
  56. Qin X.Q., Zeevaart J.A.D. (1999). The 9-cis-epoxycarotenoid cleavage reaction is the key regulatory step of abscisic acid biosynthesis in water-stressed bean. Proc. Natl. Acad. Sci. USA 96: 15354–15361 [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Saez A., Apostolova N., Gonzalez-Guzman M., Gonzalez-Garcia M.P., Nicolas C., Lorenzo O., Rodriguez P.L. (2004). Gain-of-function and loss-of-function phenotypes of the protein phosphatase 2C HAB1 reveal its role as a negative regulator of abscisic acid signalling. Plant J. 37: 354–369 [DOI] [PubMed] [Google Scholar]
  58. Schnurr J., Shockey J., Browse J. (2004). The acyl-CoA synthetase encoded by LACS2 is essential for normal cuticle development in Arabidopsis. Plant Cell 16: 629–642 [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Schwartz S.H., Qin X., Zeevaart J.A. (2003). Elucidation of the indirect pathway of abscisic acid biosynthesis by mutants, genes, and enzymes. Plant Physiol. 131: 1591–1601 [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Seo M., Koshiba T. (2002). Complex regulation of ABA biosynthesis in plants. Trends Plant Sci. 7: 41–48 [DOI] [PubMed] [Google Scholar]
  61. Seo M., Peeters A.J.M., Koiwai H., Oritani T., Marion-Poll A., Zeevaart J.A.D., Koornneef M., Kamiya Y., Koshiba T. (2000). The Arabidopsis aldehyde oxidase 3 (AAO3) gene product catalyzes the final step in abscisic acid biosynthesis in leaves. Proc. Natl. Acad. Sci. USA 97: 12908–12913 [DOI] [PMC free article] [PubMed] [Google Scholar]
  62. Sieber P., Schorderet M., Ryser U., Buchala A., Kolattukudy P., Metraux J.P., Nawrath C. (2000). Transgenic Arabidopsis plants expressing a fungal cutinase show alterations in the structure and properties of the cuticle and postgenital organ fusions. Plant Cell 12: 721–738 [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Tan B.C., Joseph L.M., Deng W.T., Liu L., Li Q.B., Cline K., McCarty D.R. (2003). Molecular characterization of the Arabidopsis 9-cis epoxycarotenoid dioxygenase gene family. Plant J. 35: 44–56 [DOI] [PubMed] [Google Scholar]
  64. Tan B.C., Schwartz S.H., Zeevaart J.A., McCarty D.R. (1997). Genetic control of abscisic acid biosynthesis in maize. Proc. Natl. Acad. Sci. USA 94: 12235–12240 [DOI] [PMC free article] [PubMed] [Google Scholar]
  65. Tang D., Simonich M.T., Innes R.W. (2007). Mutations in LACS2, a long-chain acyl-coenzyme A synthetase, enhance susceptibility to avirulent Pseudomonas syringae but confer resistance to Botrytis cinerea in Arabidopsis. Plant Physiol. 144: 1093–1103 [DOI] [PMC free article] [PubMed] [Google Scholar]
  66. Taylor I.B., Burbidge A., Thompson A.J. (2000). Control of abscisic acid synthesis. J. Exp. Bot. 51: 1563–1574 [DOI] [PubMed] [Google Scholar]
  67. Verslues P.E., Kim Y.S., Zhu J.K. (2007). Altered ABA, proline and hydrogen peroxide in an Arabidopsis glutamate:glyoxylate aminotransferase mutant. Plant Mol. Biol. 64: 205–217 [DOI] [PubMed] [Google Scholar]
  68. Wellesen K., Durst F., Pinot F., Benveniste I., Nettesheim K., Wisman E., Steiner-Lange S., Saedler H., Yephremov A. (2001). Functional analysis of the LACERATA gene of Arabidopsis provides evidence for different roles of fatty acid omega -hydroxylation in development. Proc. Natl. Acad. Sci. USA 98: 9694–9699 [DOI] [PMC free article] [PubMed] [Google Scholar]
  69. Xiao F., Goodwin S.M., Xiao Y., Sun Z., Baker D., Tang X., Jenks M.A., Zhou J.M. (2004). Arabidopsis CYP86A2 represses Pseudomonas syringae type III genes and is required for cuticle development. EMBO J. 23: 2903–2913 [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Xiong L., Ishitani M., Lee H., Zhu J.K. (2001). The Arabidopsis LOS5/ABA3 locus encodes a molybdenum cofactor sulfurase and modulates cold stress- and osmotic stress-responsive gene expression. Plant Cell 13: 2063–2083 [DOI] [PMC free article] [PubMed] [Google Scholar]
  71. Xiong L., Lee H., Ishitani M., Zhu J.K. (2002). Regulation of osmotic stress-responsive gene expression by the LOS6/ABA1 locus in Arabidopsis. J. Biol. Chem. 277: 8588–8596 [DOI] [PubMed] [Google Scholar]
  72. Xiong L., Zhu J.K. (2003). Regulation of abscisic acid biosynthesis. Plant Physiol. 133: 29–36 [DOI] [PMC free article] [PubMed] [Google Scholar]
  73. Yoshida R., Hobo T., Ichimura K., Mizoguchi T., Takahashi F., Aronso J., Ecker J.R., Shinozaki K. (2002). ABA-activated SnRK2 protein kinase is required for dehydration stress signaling in Arabidopsis. Plant Cell Physiol. 43: 1473–1483 [DOI] [PubMed] [Google Scholar]
  74. Zhu J., Lee B.H., Dellinger M., Cui X., Zhang C., Wu S., Nothnagel E.A., Zhu J.K. (2010). A cellulose synthase-like protein is required for osmotic stress tolerance in Arabidopsis. Plant J. 63: 128–140 [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Zhu J.K. (2002). Salt and drought stress signal transduction in plants. Annu. Rev. Plant Biol. 53: 247–273 [DOI] [PMC free article] [PubMed] [Google Scholar]

Articles from The Plant Cell are provided here courtesy of Oxford University Press

RESOURCES