Introduction

The outermost layer of the epidermal cell wall plays multiple roles (Carver and Gurr 2006; Leveau 2006; Pfündel et al. 2006), including the prevention of water loss from the cell surface and defense against pathogens or insects. This layer also protects plants from UV irradiation damage and prevents fusions between organs during plant development. It typically comprises the crystal epicuticular waxes deposited on the cuticle proper as well as a cuticular layer encrusted with intracuticular waxes in transit to polysaccharides (Kunst and Samuels 2003; Shepherd and Wynne Griffiths 2006).

Wax crystals are present in various shapes: tubule, fine rod, star-like rod, globule, and circular platelet (Baker 1982; Gulz 1994; Jetter and Riederer 1994; Beattie and Marcell 2002; Ao 2006). Although tubules and platelets are abundant, in rice the platelet and star-like rod types appear most frequently (Avato 1987; Yu et al. 2008). Species-unique cuticular wax components are postulated to organize themselves into crystallines and amorphous zones of the cuticle (Kirsch et al. 1997). When the concentration of a certain component reaches a critical threshold, epicuticular crystalloids can arise, on the aerial surface, from the amorphous wax mixture that is embedded in the cutin polymer (Jetter and Riederer 1994; Jeffree 1996).

Most wax mutants, such as eceriferum1 (cer1), cer2, cer3, cer4, cer5, cer6, glossy1 (gl1), gl8, and wax deficient anther1 (wda1), have less crystal wax on their cell surfaces (Lemieux 1996; Hansen et al. 1997; Xu et al. 1997; Jung et al. 2006). However, some show an altered epicuticular wax structure. For example, knb1 mutants contain smaller flake-like wax crystals and bcf1 mutants possess smaller plate-like crystals as well as long, slender, ribbon-like crystals (Jenks et al. 1996). In gl1 and gl26 mutants, their crystal shapes are changed from platelet-type to globule-type (Beattie and Marcell 2002). gl1 also manifests a pleotropic effect, e.g., less wax, small trichomes, and a thin cuticle membrane structure. GL1 is a membrane-bounded desaturase/hydroxylase (Sturaro et al. 2005).

Cutin is composed of inter-esterified hydroxyl and hydroxyl epoxy fatty acids derived from common cellular fatty acids, mostly C16–C18 fatty acids with ω- and mid-chain hydroxyl groups (Kolattukudy 2001). However, α, ω-dicarboxylic fatty acids are the major cutin monomers in Arabidopsis, where ω-hydroxy and mid-chain hydroxylated fatty acids are also found (Bonaventure et al. 2004; Franke et al. 2005). Although the predominant cutin monomers are known, some unidentified components in cutin polymer and structure are poorly understood. In addition, mutants with a defective cutin matrix and/or inadequate organization are rare. Although the synthesis of cutin must be coordinated with extension of the cell surface during growth, how this is achieved also remains unsolved.

A defect in LACERATA (LCR) encoding a cytochrome P450 of the CYP86 family is implicated in the epidermal biosynthesis of cutin, causing cutin deformation and organ fusion (Wellesen et al. 2001). Aberrant induction of type three genes 1 (ATT1) encodes a cytochrome P450 monooxygenase, catalyzing fatty acid oxidation for the biosynthesis of extra-cuticular lipids. In att1, cutin content is reduced to 30% and the cuticle is loosely organized, thereby increasing the transpiration rate two-fold (Xiao et al. 2004). The eibi1 mutant is essential for drought resistance in barley. Normally, EIBI1 functions in the cuticle as the transport-limiting layer; eibi1 has the highest relative rate of water loss among the known wilt mutants. Leaves of eibi1 have high stomatal density and a great chlorophyll efflux in 80% ethanol (Chen et al. 2004). Sorghum bicolor bloomless22 (bm22) mutants also show a reduced cuticle thickness and increased water loss (Jenks et al. 1994). In contrast, the amount of cutin is enhanced in the bodyguard (bdg) mutant even though the cuticle is loosely organized (Kurdyukov et al. 2006). There, mutant plants exhibit strong growth retardation and abnormal leaf morphology. When immersed in 80% ethanol, leaves of bdg release chlorophyll faster than do wild-type leaves.

Here, we describe the identification of a rice mutant defective in its organization of cuticle and wax crystals, which leads to increased water loss via diffusion from the leaf surface.

Materials and methods

Plant materials and growing conditions

Generation of T-DNA tagging rice plants (Oryza sativa ssp. japonica cv. Dongjin and Hwayoung) has been reported previously (Jeon et al. 2000; Jeong et al. 2002). Seeds were germinated on a 1/2 MSO medium solidified with 0.2% phytagel and 0.55 mM myo-inositol (Sigma–Aldrich) in a closed container. The seedlings were grown for 1 or 2 weeks at 30°C under continuous light. They were then transplanted to soil in the greenhouse or paddy field for an additional 2 weeks of development.

Isolation of wdl1 knockout plants

Three wdl1 knockout alleles were isolated from the rice-flanking sequence database (http:://www.postech.ac.kr/life/pfg) (An et al. 2005a, b). T2 progeny of the primary insertional mutants were grown to maturity for seed amplification. Genotypes of wdl1-1 were determined by PCR with the following primers: F1 (5′-ACCAGATCAAGTGGAGTGGA-3′), R1 (5′-AAAGGCAACAGTCAAGCAAG-3′), and LB-1 (5′-CATCTTGAAC GATAGCCTTT-3′). For genotyping wdl1-2, primers F2 (5′-CTCGCATCTTGATTCCTCAT-3′), R1, and LB-1 were used; for wdl1-3, F3 (5′-CGTGCAACCTTCTTTGGTAA-3′), R2 (5′-CCATTGGATAGTCCTGTGGA-3′), and RB-1 (5′-ATCCAGACTGAATGCCCACAGGC-3′).

Quantitative real-time PCR

Expression patterns for WDL1 were measured by real-time PCR with a Roche LightCycler II. UBQ mRNA expression was used to normalize the expression ratio for a gene. Primers included 5′-GATGGACCTGAAGGTGGTT-3′ and 5′-AACCACCTTCAGGTCCATC-3′ for WDL1, 5′-GGTGCTCAACACATCCACT-3′ and 5′-ATGCATTCTCTGCTCTCCT-3′ for ABA2, and 5′-CACGGTTCAACAACATCCAG-3′ and 5′-TGAAGACCCTGACTGGGAAG-3′ for UBQ. Changes in gene expression were calculated via the ∆∆Ct method.

Analyses by scanning electron microscopy and transmission electron microscopy

The second leaves from 7-day-old seedlings grown in a closed container were used for live SEM, and for SEM and TEM analyses. For live SEM, fresh samples were sputter-coated directly with palladium and examined with a scanning electron microscope (LEO 1450VP; Carl Zeiss, Jena, Germany). Other samples for SEM and TEM were pre-fixed for 3 h with 3% glutaraldehyde–sodium phosphate buffer (0.1 M) at room temperature and rinsed three times with 0.1 M sodium phosphate buffer. Post-fixation was performed with 2% OsO4 at 4°C. The samples were dehydrated through an ethanol series and infiltrated with an isoamyl acetate series. TEM samples were sectioned 40–60 nm thick and then stained with 2.5% uranyl acetate before examination with a transmission electron microscope (JEOL 100CX-I, 80 kV; JEOL, Japan).

Analysis of leaf waxes and cutin

Before extracting wax, we scanned the leaves to calculate their surface areas. For total extraction, leaf blades and sheaths were excised from seedlings at 14 days after germination (DAG) and immersed for 1 h in 20 ml of CHCl3 at 55°C (Haas et al. 2001). To avoid removing membrane lipids, the severing points of leaves were not immersed. As an internal standard, 20 μg of n-tetracosane (Fluka) was added to each sample. Chloroform was evaporated under a gentle stream of N2 to a final volume of 100 μl. The wax samples were derivatized with 20 μl of pyridine and 20 μl of BSTFA (Machery-Nagel, Düren, Germany), then analyzed by GC–MS as previously described (Jung et al. 2006; Yu et al. 2008). Quantitative determination of wax components was based on the equivalent ratio of mass to peak area between the component and internal standard. Wax monomers were identified by mass spectrometry (Agilent Technologies, Böblingen, Germany). After this extraction, the same leaves were used in cutin analysis performed as described by Franke et al. (2005).

Determination of transpiration and conductance

Transpiration rates were measured with a steady-state porometer (LI-1600; LI-COR Bioscience, Lincoln, NE, USA) using 50 DAG plants grown under greenhouse conditions (Koizumi et al. 2007; Woo et al. 2007). The clip of a porometer evaluated water movement for each sampled leaf for 1 s and then recorded the value. Day and night evaporation rates were measured at 2 p.m. and 11 p.m.

Cell numbers and leaf fresh weights

Aerial parts from plants grown for 7 days in a closed container were held at 30°C and 40% relative humidity. For each genotype, the fresh weights of excised leaves were measured for water loss at 0, 5, 10, 20, 30, 60, 120, and 180 min. To count the number of stomatal and pavement cells per unit area, we attached the second leaves from 7 DAG seedlings to adhesive tape. Their epidermal and mesophyll cells were then removed through washing, leaving an epidermal cell trace on that tape. Densities of the stomatal and epidermal pavement cells, as well as the stomatal index of adaxial surfaces, were determined according to the procedure of Gray et al. (2000).

Chlorophyll leaching assay

After 7-day-old seedlings were air-dried for 6 h, leaf samples were incubated for 1 h in 80% ethanol at 37°C. The amount of extracted chlorophyll was measured at A647 and A664 nm on a UV-1,700 Pharmaspec spectrophotometer (Shimadzu). Chlorophyll concentrations were calculated as described previously (Lolle et al. 1997).

Vector construction and sub-cellular localization

The coding region of WDL1 cDNA was PCR-amplified with primers 5′- CCCGGGATGCTTGGTTTTGCGCCG-3′ (the ATG start codon of OsWDL1 is underlined) and 5′-ACTAGTGTCCCATTGGATAGTCCTGT-3′ (the TAG stop codon was deleted to make a reading-frame fusion between WDL1 and sGFP). The fragment was cloned into the SmaI and SpeI sites between the maize Ubiquitin promoter and the sGFP coding sequence (Moon et al. 2008). A vector containing the AtBIP–mRFP fusion was obtained from Inhwan Hwang (Park et al. 2004). Protoplasts were produced from rice mesophyll cells as follows. Leaves of 7 DAG seedlings were diced with a razor blade and incubated in a cell wall digestion-enzyme solution containing 1.5% cellulase, 0.3% macerozyme, 0.1% pectolyase, 0.6 M mannitol, 10 mM MES, 1 mM CaCl2, and 0.1% (w/v) bovine serum albumin for 4 h at 26°C with gentle agitation (75 rpm), as described previously (Moon et al. 2008). Protoplasts (1 × 106 per ml) were mixed with 10 μg each of WDL1-sGFP and AtBIP-mRFP DNAs, then electroporated in 4-mm-wide cuvettes with a Gene Pulser Xcell (BioRAD, Baltimore, MD, USA) that was set at 300 V and 450 μF. For transient expression in onion cells, the epidermis was bombarded via the biolistic particle delivery system (PDS−1,000, Dupont), with parameters of 0.4 mg per shot of gold beads coated with about 2 μg per shot of each plasmid DNA and 900 psi of pressure in a 25-in. Hg vacuum. After 10 to 15 h of incubation at 25°C under darkness, the transformed protoplasts and onion epidermal cells were examined on a Zeiss Axioplan fluorescence microscope equipped with filter sets for GFP and RFP.

Results

Isolation of the wdl1 mutant from T-DNA tagged lines

We isolated a conditional lethal mutant line, 3A-07662 (cv. Dongjin), from a T-DNA insertional mutant population in japonica rice (Jeon et al. 2000; An et al. 2003; Lee et al. 2003; Ryu et al. 2004). These mutant plants were recessive-lethal in the paddy field. Mutant seeds produced from heterozygous plants germinated normally, but the seedlings showed a severe growth retardation phenotype and died within 30 days. However, when cultured under high humidity (more than 90% relative humidity), plants grew for an extended period although their stems and leaves were shorter than normal (Fig. 1a, b). Under low humidity, i.e., 40%, leaves of those mutants wilted within 10 min (Fig. 1c, d). Most mutant plants grown under normal greenhouse conditions died (Fig. 1e) but some occasionally survived to maturity and produced sterile spikelets (Fig. 1f). These survivors were shorter and their leaves fragile compared with their segregating wild-type siblings (Fig. 1f). Such phenotypes appeared to arise from sensitivity to dehydration. We named this mutant wilted dwarf and lethal 1 (wdl1).

Fig. 1
figure 1

Phenotypes of rice seedlings. a Heights of WT (left), wdl1-1 (2nd from left), wdl1-2 (3rd from left), and wdl1-3 (right) grown in closed container for 7 days after germination (DAG). b Third leaves of 7 DAG plants in same order as a. c Detached leaves were dried for 20 min at room temperature. d Enlargement of c at tip region. e Plants grown for 7 additional days in soil under natural conditions. Left, WT; right, wdl1-1. f WT (left) and wdl1-1 (right) plants at mature stage

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T-DNA insertional mutations in WLD1

Sequence analysis of the T-DNA flanking region from the mutant revealed that T-DNA was inserted into LOC_Os11g48070 (http://tigrblast.tigr.org/euk-blast/) on Chromosome 11. Its full-length cDNA was identified as AK067429 in the Knowledge-based Oryza Molecular Biological Encyclopedia (KOME) (http://cdna01.dna.affrc.go.jp/cDNA). The primary structure of WDL1 comprises six exons and five introns (Fig. 2a). In wdl1-1, T-DNA is located 1,203 bp downstream from the ATG start codon, in the second intron of the gene.

Fig. 2
figure 2

Isolation of wdl1 knockout plants. a Schematic diagram of WDL1 and insertion positions of T-DNA. Closed boxes represent 6 exons; connecting black lines are 5 introns. Horizontal arrows indicate primers used for genotyping T2 progeny. b Alignment of amino acid sequences of SGNH proteins. Black boxes represent identical amino acids; gray boxes, similar amino acids. Blocks are conserved domains; dots are active amino acids in SGNH subfamily. c Expression analysis of WDL1 in WT and three wdl1 alleles by quantitative real-time PCR. Y axis, relative values between transcript levels of WDL1 and UBQ; vertical bars, standard deviation

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The predicted protein encoded by WDL1 contains 260 amino acid residues, with a molecular mass of approximately 28 kDa. This protein is 63% identical to cowpea CPRD49 (CowPea clones Responsive to Dehydration 49, BAB33036.1), which is induced by dehydration (Iuchi et al. 1996). WDL1 is also 67% identical to At3g11210 (AAM63310.1) and 51% to At2g38180 (AAL31218.1). Functional-domain analysis with Pfam 7.0 (http://www.sanger.ac.uk/Software/Pfam) indicated that the region between the 13th and 207th amino acid residues is highly conserved among these proteins (Fig. 2b). This region contains the conserved motifs found in SGNH subfamily proteins, which are members of the GDSL lipase superfamily (Akoh et al. 2004). SGNH proteins have four invariant catalytic residues—Ser, Gly, Asp, and His—in blocks I, II, III, and V, respectively (Fig. 2b; Akoh et al. 2004). WDL1 contains these conserved residues at Ser19, Gly48, Asp83, and His194. Phylogenic analysis using the MEGA 2.0 program indicated that the WDL1 protein is distantly related to other members of the GDSL lipase gene family (Fig. S1). From the flanking sequence database of T-DNA insertion sites (An et al. 2003; Jeong et al. 2006), we identified two additional alleles: wdl1-2 (in the first intron) and wdl1-3 (in the fifth exon) from Lines 4A-01860 (cv. Dongjin) and 1B-07129 (cv. Hwayoung), respectively (Fig. 2a). Quantitative RT PCR analyses of RNA extracted from homozygous mutant plants indicated that those three were null alleles (Fig. 2c). All had the same wilted dwarf phenotype and most died as seedlings. Therefore, we concluded that these phenotypes are due to the disruption of WDL1 expression.

Because our insertional lines showed conditional mutant phenotypes, we examined whether gene expression is inducible. Although not influenced by all types of stress, transcript levels were reduced by wounding, drought, salt, and ABA treatments (Lee et al. 2006; Fig. 3a). As a control we also measured ABA2 transcript levels during these trials (Rabbani et al. 2003); as expected, those were increased by such treatments (Fig. 3b), indicating that our stresses were properly applied. Although WDL1 was expressed in whole plants, transcript levels were higher in the shoots and young panicles than in the calli and roots (Fig. 3c). Levels were also high in the reproductive organs.

Fig. 3
figure 3

WDL1 gene expression pattern. aWDL1 transcript levels after treatment by wounding, drought, 250 mM NaCl, or 100 μM ABA for 30 min, 1 h, or 2 h. bABA2 expression after same treatments as a for 30 min, 1 h, or 2 h. cWDL1 expression patterns in callus, root at 7 DAG, shoot at 7 DAG, panicle <8 cm, mature leaf at 80 DAG, mature flower, and seed at 5 days after pollination. Y axis, relative WDL1 transcript level; vertical bar, standard deviation

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wdl1 mutations affect epidermal permeability

Because wdl1 mutants wilted under low humidity, we examined the surface structure of their leaf blades and found that the stomatal and pavement cells were smaller than in the wild type (WT) (Fig. 4a–d). This decrease in size was also associated with a higher number of stomatal and pavement cells per unit area. The ratio between the two cell types was increased by about 12% in the mutant (Fig. 4e).

Fig. 4
figure 4

Analyses of stomatal cell (SC) and pavement cell (PC) densities. a, b SEM and c, d optical microscopy of cells in adaxial region of 2nd leaves from a, c WT and b, d wdl1-1 plants. Scale bars are 50 μm. e SC density, PC density, and stomatal index in adaxial region of 2nd leaves. For each genotype, 24 samples were measured from 5 leaves. Stomatal index is number of stomata as percentage of total cells

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The transpiration rates for mutant plants were 2.3 times higher during the day compared with their WT siblings (Fig. 5a, b). Chlorophyll leaching assays are used for measuring the non-stomatal permeability that is presumably indicative of water loss (Yoshida et al. 2002). To minimize the water loss through the stomata during our assay, we dried the leaf samples for 6 h before treating with 80% ethanol. It was previously reported that detached leaves close their stomata within 30 min (Kerstiens et al. 2006). Chlorophylls were extracted three- to four-fold more rapidly from all three wdl1 seedlings relative to the WT (Fig. 5c, d). All mutants lost water more quickly over the first 30 min (Fig. 5e). These data indicated that conductance is faster in wdl1, probably due to non-stomatal diffusion through their leaf surfaces.

Fig. 5
figure 5

Physiological characterization of wdl1 mutants. Water evaporation rates were measured at 2 p.m. (a) and 11 p.m. (b). c Chlorophyll extraction for 60 min. d Chlorophyll leaching from WT and mutants. e Percentage of fresh weight reduction in detached leaves under 40% relative humidity at 30°C. Experiments were repeated 3 times. Each bar indicates mean ± SD from 3 replicates

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The cuticle is abnormal in wdl1 mutants

SEM analyses showed that the WT leaf surface was covered evenly with hairy crystal waxes (Fig. 6a–c). This is consistent with previous reports (Sánchez et al. 2003; Jung et al. 2006; Yu et al. 2008). In contrast, the leaf surfaces of wdl1 mutants grown under the same conditions of high humidity were covered by both normal and disorganized crystal waxes, including a split wax cluster, curdled wax, and a stretched wax curd (Fig. 6e–g, i–k). To confirm that these changes were not due to any alteration during sample preparation we examined the surfaces with a live SEM. Our analyses presented the same results, i.e., abnormal waxes on the mutant surface (Fig. 6c, g).To examine whether the substances from those mutant and WT leaves are indeed waxes, we washed the samples with 100% chloroform (Haas et al. 2001). Images obtained after this washing demonstrated that the surfaces were cleaned in both types (Fig. 6d, h). These results demonstrated that mutations in WDL1 affect wax crystallization and distribution on the leaf surface. Cross-sectioning of the WT showed that the cuticle was even, flat, and regular (Fig. 7a, f) whereas the wdl1 cuticle was loosely packed and had unclear boundaries, uneven thickness, and less compactness (Fig. 7b–e, g–j). Cell walls of the WT were of even and regular thickness (Fig. 7a) compared with an irregular and fluctuating thickness from the mutant (Fig. 7b–e).

Fig. 6
figure 6

Scanning electron microscopy analyses of adaxial leaf surfaces from WT, wdl1-1, wdl1-2, and wdl1-3. a–c WT showing star-type wax crystals. e–g,i SEM of wdl1-1. Live SEM for WT (c) and wdl1-1(g). SEM for WT (d) and wdl1-1(h) after wax was removed by chloroform. j wdl1-2, and k wdl1-3. f Star-type and split-wax cluster, i curdled wax structure, j stretched curd wax and. P, papillae; SC, stomata cell. Scale bars are 10 μm (a, d) and 2.5 μm (c, e–k)

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Fig. 7
figure 7

Transmission electron microscopy analyses of adaxial leaf epidermis from 7-day-old seedlings. (a, f) WT showing even thickness, regular compactness, and clear cuticle boundary. (b–e, g–h) TEM of wdl1-1 with uneven thickness, less compactness, and vague cuticle boundary (g), even but occasionally thin and irregular region (h). i TEM of wdl1-2 and j wdl1-3. CM, cuticle membrane; CW, cell wall

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WDL1 protein localizes in the ER

Because the wdl1 cuticle was abnormal, we hypothesized that the WDL1 protein is involved in wax and cutin formation. If true, that protein would likely be localized to the endoplasmic reticulum, where most wax and cutin biosynthesis occurs. Therefore, we made a fusion of the synthetic GREEN FLUORESCENCE PROTEIN (sGFP) at the C-terminal of WDL1 (Fig. 8a, upper). As a positive control, we used the previously characterized AtBIP–mRFP that is localized to the ER (Fig. 8a, lower; Park et al. 2004). When these chimeric molecules were co-introduced into rice mesophyll protoplasts prepared from 10-day-old seedlings, the GFP signal from WDL1–GFP coincided with the RFP signal driven by the AtBIP–mRFP protein (Fig. 8b–e). As a negative control, we used sGFP localized to the nucleus and cytoplasm (Fig. 8f–h). Similar results were obtained when these fusion molecules were introduced into onion epidermal cells (Fig. 8i–k), thereby indicating that WDL1 is an ER protein and may be related to wax and cutin biogenesis.

Fig. 8
figure 8

Sub-cellular localization of WDL1–GFP. a Schematic diagrams of 35S:AtBIP–RFP and Ubi:WDL1–GFP. (b–e) Sub-cellular localization of WDL1 in mesophyll cells. b Localization of WDL1–GFP protein and c AtBIP–RFP as ER marker. d Auto-fluorescence of chloroplast. e Merged image of b, c, and d. (f–h) Sub-cellular localization of sGFP in mesophyll cells. f sGFP image, g auto-fluorescence, and merged image of f and g h. (i-k) Sub-cellular localization of WDL1::GFP in onion epidermal cells. i WDL1–GFP, j AtBIP–RFP, and k merged images of h and i

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Amounts of wax and cutin are not significantly changed in wdl1

Leaves of wdl1 had a disorganized epicuticular crystal wax layer and deformed epidermal cell arrangement. To determine whether this abnormality is due to a change in wax composition in the cuticle, we sampled leaves from WT and wdl1 plants grown for 14 days under high humidity. Epicuticular waxes were extracted with chloroform and analyzed by GC–MS. The total amounts of wax did not differ significantly (Fig. 9a), and detailed analyses showed that the amounts of fatty acids, aldehydes, alkanes, and esters also remained the same. However, the content of primary alcohol monomers was slightly increased in the mutant (Fig. 9b), a change mainly due to a rise in C30 alcohol (Fig. 9c).

Fig. 9
figure 9

Epicuticular wax and cutin comparative analyses. a Total wax in epicuticular region from WT, wdl1-1, and wdl1-2. Total waxes were prepared from 14 DAG seedlings grown under high humidity. b Wax monomers from WT, wdl1-1, and wdl1-2. Fatty acids (FA), Aldehydes (Ah), Primary Alcohol (PA), Alkanes (Ak), Esters (E). c Primary alcohols from WT, wdl1-1, and wdl1-2. d Total cutin amounts in cuticle layers from WT and wdl1-1. e Amounts of cutin monomers from WT and wdl1-1. Each bar indicates mean ± SD from 4 replicates; experiments were repeated 3 times

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We also analyzed cutin that remained after the epicuticular wax was extracted. This represented the lipid polymers, e.g., cutin and other covalently bound lipids, attached to the cell wall (Franke et al. 2005; Kurdyukov et al. 2006). Here, the mutants and wild type did not differ significantly in their amounts of cutin (Fig. 9d, e).

Discussion

Wilt phenotype is associated with increased permeability

We have now identified T-DNA insertional mutants that are dwarf and lethal when grown under normal humidity. There phenotypes are associated with a higher day time transpiration rate. Our chlorophyll leaching assay suggested increased permeability of the wdl1 leaf surface. Although the contribution of stomatal and non-stomatal water loss during the day has not yet been determined, our results suggest that wdl1 mutants might lose water more rapidly, at least partially because of enhanced cuticle-membrane permeability. TEM examination showed that the cuticle of wdl1 is loose and irregular, which likely accounts for this greater permeability. The wdl1 cuticle membrane is also thicker and uneven, with such morphological changes probably leading to enhanced conductance.

Although some researchers have reported no clear relationship between cuticle water permeability and physical properties, e.g., thickness, coverage, and uniformity (Riederer and Schreiber 2001; Kerstiens 2006), analysis of Sorghum bicolor bloomless (bm) mutants with an altered epicuticular wax structure has identified a mutation affecting the deposition of both epicuticular wax and cuticle (Jenks et al. 1994). The cuticle from bm plants is about 60% thinner and only 20% of the WT weight. This diminished cuticle deposition is correlated with increased epidermal conductance. Similar observations have been made with the Arabidopsis wax2 mutant, in which the cuticle is thicker and structurally disorganized (Chen et al. 2003). Lacs2 (Long-Chain Acyl–CoA Synthetase 2) has a thinner abaxial cutin and an increased rate of chlorophyll leaching (Schnurr et al. 2004). The att1 mutant, in which cutin content is reduced to 30%, also has a loose cuticle and greater permeability to water vapor (Xiao et al. 2004).

Because our mutants also presented a loosely packed and uneven cuticle, we should have expected a change in the cutin amount but, instead, found no significant alteration in total cutin and cutin monomer levels. Therefore, WDL1 does not appear to be involved in cutin monomer biosynthesis, making it probable that the gene product affects cutin polymer organization. Nevertheless, it is still unknown what determines this organization. In att1 and bm mutants, the cuticles are loose due to a lower cutin content that results in more rapid transpiration (Xiao et al. 2004). This would suggest that reducing cutin levels affects the structure of the cuticle. However, a higher level of cutin in bdg mutants also causes the cuticle proper to be less dense, implying that compactness is not always correlated with cutin amount (Kurdyukov et al. 2006). Therefore, other factors must control this organization. Because our wdl1 mutants have an altered structure in their cuticle membrane but little change in the level of cutin, we might conclude that the WDL1 gene product is involved in cutin organization. This finding is similar to that reported for the wax2 mutant, where no effect is found on cutin amount even though plants have a thicker and less opaque cuticle proper (Rowland et al. 2007). Cutan, a non-depolymerizable or unsaponifiable polymethylene biopolymer that is associated with the cuticle membrane (Schreiber 2005), plays an important role as a barrier to the sorption of polar and non-polar substances, as well as other aromatic compounds (Stimler et al. 2006). Because we did not compare the amount, composition, and organization or arrangement of cutan between mutant and wild type, we cannot exclude the possibility that different cutan levels can account for the differences in transpirational and chlorophyll leaching rates in the wdl1 mutant. Alternatively, WDL1 could indirectly affect the cuticle. For example, although SHN gene is not involved in cutin biosynthesis, overexpression of the gene exerted influence in deforming cuticle structure and increased cuticle permeability (Aharoni et al. 2004). The gene is a member of AP2/EREBP transcription factors that activates terpenoid biosynthesis.

Abnormal wax crystals are likely due to an irregular cuticle

Because wdl1 mutants have an altered epicuticular wax structure, we predicted a significant change in the amount and composition of wax compared with the WT. However, the levels of fatty acids, aldehydes, alkanes, and esters remained constant, while only a slight increase was observed in the primary alcohols, especially the C30 chain length. Because these primary alcohols are the major component of waxes, it is possible that a rise in their levels modifies the overall organization of those waxes. For example, a 1.6-fold increase in alkanes and primary alcohols in knb1 accompanies a change in the wax crystal structure (Jenks et al. 1996) while a decline in wax monomer levels also affects the epicuticular wax structure. In cer4 mutants, the levels of C24, C26, C28, and C30 primary alcohols are severely reduced, and the wax crystal structure shifts to a vertically oriented plate-like shape (Rowland et al. 2006).

When the amount of wax exceeds a critical threshold, crystals are generated from the wax mixture that is embedded in the crosslinked cutin (Jetter and Riederer 1994; Jeffree 1996; Kirsch et al. 1997). Because of the loose, uneven, and variable arrangement in the cuticle membrane and cell walls of our wdl1 mutants, it is possible that wax is irregularly secreted, thereby leading to disorganized deposition.

Defects in the cuticle are not compensated for by a greater amount of epicuticular wax. For example, in bdg mutants with a 3.5-fold increase in wax content, chlorophyll is extracted twice as fast as from the wild type (Kurdyukov et al. 2006). Furthermore, although shn has six-fold more wax per unit area compared with the WT, its rates of chlorophyll and water extraction are eight-fold and six-fold faster, respectively (Aharoni et al. 2004).

WDL1 is likely involved in a later step of cutin formation

WDL1 is homologous to SGNH subfamily proteins, which are members of the GDSL lipase superfamily (Akoh et al. 2004). That superfamily comprises a diverse group of hydrolases, including lipases, esterases, thioesterases, arylesterases, proteases, and lysophospholipases. These enzymes display various functional properties, such as broad substrate specificity and region specificity (Akoh et al. 2004). Compared with common lipases, the GDSL lipases have a flexible active site that changes conformation in the presence of substrate. WDL1 has a catalytic triad consisting of the Ser19, Asp191, and His194 residues in the active center of the SGNH family proteins (Ling et al. 2006). A group of SGNH lipases has been characterized as sinapine esterase (Clauβ et al. 2008). In addition to sinapine esterase activity, the SGNH lipases show broad substrate specificity toward various choline esters, including phosphatidylcholine. To investigate whether WDL1 is a lipase, we examined acyl-hydrolase activity of heterologously expressed protein, using 4-nitrophenyl derivatives as substrates. Although we were unable to demonstrate hydrolase activity for WDL1 using this model substrate, the enzyme probably functions as a lipase because the protein shares high sequence homology with SGNH lipases and contains conserved residues in the active center of the SGNH proteins (Ling et al. 2006). However, either their very narrow substrate specificity hampers functional analysis or else this enzyme functions as part of a multi-complex system. We also cannot discount the possibility that WDL1 is active in the formation of ester linkages involving non-lipid cell wall components such as methylated and acetylated carbohydrates. Secondly, these could affect the anchorage of cutin polyester that disturbs the cuticular membrane.

WDL1 is co-localized with an ER marker protein, indicating that it is a protein within the endoplasmic reticulum. Because most wax and cutin monomers are synthesized there, results from our localization experiment demonstrate that WDL1 protein also plays a role either in providing or connecting and arranging the precursors for proper cuticle formation. Because our biochemical analyses did not reveal any significant alteration in the monomers, it is likely that WDL1 is involved in steps of cutin organization, perhaps by providing pre-formed oligomeric esters or inducing the modification of non-ester cross-links, e.g., C–C and C–O–C, of non-depolymerizable components.