Maize glossy6 is involved in cuticular wax deposition and drought tolerance

glossy6, a novel maize gene, is involved in the intracellular trafficking of waxes, opening the door to studying the necessary yet poorly understood process of wax loading on plant surfaces.

Abstract

Cuticular waxes, long-chain hydrocarbon compounds, form the outermost layer of plant surfaces in most terrestrial plants. The presence of cuticular waxes protects plants from water loss and other environmental stresses. Cloning and characterization of genes involved in the regulation, biosynthesis, and extracellular transport of cuticular waxes onto the surface of epidermal cells have revealed the molecular basis of cuticular wax accumulation. However, intracellular trafficking of synthesized waxes to the plasma membrane for cellular secretion is poorly understood. Here, we characterized a maize glossy (gl6) mutant that exhibited decreased epicuticular wax load, increased cuticle permeability, and reduced seedling drought tolerance relative to wild-type. We combined an RNA-sequencing-based mapping approach (BSR-Seq) and chromosome walking to identify the gl6 candidate gene, which was confirmed via the analysis of multiple independent mutant alleles. The gl6 gene represents a novel maize glossy gene containing a conserved, but uncharacterized, DUF538 domain. This study suggests that the GL6 protein may be involved in the intracellular trafficking of cuticular waxes, opening the door to elucidating the poorly understood process by which cuticular wax is transported from its site of biosynthesis to the plasma membrane.

Introduction

The hydrophobic cuticle covers most aerial parts of land plants and acts as a barrier to protect plants from non-stomatal water loss, ultraviolet light, physical damage caused by insects or fungi, and other biotic or abiotic stresses (Shepherd and Wynne Griffiths, 2006). The cuticle mainly consists of two types of lipophilic material, cutin and cuticular wax. Cutin is the major structural component of the cuticle and is composed of hydroxy and epoxy C₁₆ and C₁₈ fatty acid monomers, as well as glycerol (Nawrath, 2006). Cuticular waxes, as the second component of the cuticle, are either interspersed in the cutin matrix (intracuticular waxes) or overlie the outermost surface of cutin polymer (epicuticular waxes). Cuticular waxes consist of complex mixtures of hydrophobic compounds, mostly very-long-chain fatty acids (VLCFAs) with more than 20 carbon atoms and their derivatives, including primary and secondary alcohols, aldehydes, alkanes, ketones, and wax esters (Lemieux, 1996; Kunst and Samuels, 2003; Samuels et al., 2008). Cuticular wax composition and amounts vary greatly among plant species, tissues and organs, and development states (Lee and Suh, 2015). Primary alcohols and aldehydes are the major components of the cuticular waxes in juvenile maize leaves (Javelle et al., 2010).

Cuticular wax biosynthetic pathways have been extensively studied in Arabidopsis by identification and functional characterization of wax-deficient mutant genes (Lee and Suh, 2013). The first step in wax biosynthesis is the elongation of C₁₆ and C₁₈ fatty acids in the endoplasmic reticulum (ER) into VLCFAs by joining C₂ building blocks of acetyl coenzyme A into a straight-chain of up to 34 carbon atoms via a fatty acid elongase complex (Samuels et al., 2008; Kunst and Samuels, 2009). Following elongation, VLCFAs are modified into various wax products via the distinct alcohol-forming and alkane-forming pathways (Bernard and Joubès, 2013).

Secretion of cuticular waxes was elucidated via the identification of two membrane-located ATP binding cassette (ABC) transporters, CER5 and WBC11, responsible for wax export across the plasma membrane (PM) (Pighin et al., 2004; Bird et al., 2007). However, the mechanism for intracellular trafficking of wax components from their site of synthesis at the ER to the PM is less clear. Through the characterization of the Arabidopsis ltpg mutant, lipid transfer proteins have been proposed to be involved in cuticular wax deposition (Debono et al., 2009). In addition, deficient wax secretion in mutants of the Arabidopsis GNL1 or ECH genes that function in endomembrane vesicle trafficking indicated that altered ER morphology impacts wax biosynthetic capacity (McFarlane et al., 2014).

In maize (Zea mays), our understanding of cuticular wax biosynthesis and accumulation has been aided by the identification of more than 30 glossy (gl) loci (Schnable et al., 1994; and unpublished data from the Schnable lab), some of which have been cloned. gl1 and gl2, homologs of Arabidopsis CER3/WAX2 and CER2, are involved in the leaf cuticular wax alkane-forming pathway and the extension of VLCFAs to C₃₀ (Tacke et al., 1995; Negruk et al., 1996; Hansen et al., 1997; Sturaro et al., 2005). gl4 and gl8, homologs of KSC6 and KCR of Arabidopsis, belong to the fatty acid elongase complex and play important roles in VLCFA synthesis (Xu et al., 1997; Dietrich et al., 2005; Liu et al., 2009). gl3 and gl15 encode MYB and APETALA2 (AP2)-like transcription factors respectively, and both function in the regulation of cuticular wax biosynthesis (Moose and Sisco, 1996; Liu et al., 2012). The gl13 gene encodes a putative ABC transporter involved in the transport of epicuticular waxes (Li et al., 2013).

One of the most important functions of plant cuticular waxes is to serve as a protective barrier against environmental stresses, including drought. Drought stress alters the composition and increases the content of cuticular waxes in Arabidopsis, rice, and wheat in some cases, and cuticular wax content has been associated with drought tolerance (Aharoni et al., 2004; Kosma et al., 2009; Zhu and Xiong, 2013; Zhang et al., 2015). Some genes involved in biosynthesis and transport of cuticular wax may be used to improve plant drought tolerance in Arabidopsis and several crops (Lee and Suh, 2015; Xue et al., 2017). However, the role of cuticular wax accumulation in drought tolerance in maize remains unclear.

Here, we report the cloning of the maize glossy6 (gl6) gene that is involved in cuticular wax accumulation and show that the gl6 mutant, relative to wild-type, exhibits reduced epicuticular wax accumulation, as well as increased cuticle permeability and seedling drought sensitivity. Further study suggests that the GL6 protein may be involved in intracellular transport of cuticular wax, providing novel insight into the cuticular wax biosynthesis/transport pathway.

Materials and methods

Plant material

The maize (Zea mays) glossy6 reference mutant allele (termed gl6-ref, Schnable lab Ac#245) was obtained from the Maize Genetics Stock Center and maintained in the Schnable lab (Schnable et al., 1994). Four additional glossy6 alleles (gl6-2, gl6-3, gl6-4, and gl6-5) isolated via Mu transposon tagging screens in the Schnable lab from 1992 to 2010 were used to clone the gl6 gene via Seq-Walking and digestion–ligation–amplification (DLA) analysis. Three ethyl methanesulfonate (EMS)-induced alleles (gl6-6 and gl6-7 were generated by the Schnable lab; gl6-8 was generated by M. G. Neuffer’s lab, GN) were also screened and used to verify the candidate gl6 gene in this study (Supplementary Table S1 at JXB online; Schnable et al., 1994).

The gl6-ref mutant has been backcrossed to B73 inbred line background up to eight generations and then continuously self-pollinated. BC₈F₂ segregating populations from this backcrossing program were used to map the gl6 gene by the BSR-Seq technology, and subsequent BC₈F₃ homozygous gl6 mutant or wild-type lines were used for the drought tolerance assay, physiology characterization, and leaf surface wax analyses.

Electron microscopy techniques

The second leaves collected from gl6-ref mutant and wild-type were used for field emission scanning electron microscopy (FE-SEM) analysis according to standard protocols (Aharoni et al., 2004). In brief, samples were fixed on the spindle and frozen in liquid nitrogen, dried in a vacuum-drying oven and then characterized with a FE scanning electron microscope (SU8010, Hitachi, Japan).

The collected leaves were also used for transition electron microscopy (TEM) analysis by the conventional chemical protocols (Chen et al., 2003). Samples were fixed in 2% glutaraldehyde in 0.1 M phosphate buffer (pH 7.4), post-fixed in 1% osmium tetroxide for 1 h, dehydrated with gradient alcohol solutions, and embedded in LR White resin. Ultrathin cross-sections were prepared with a Leica EM UC6 ultramicrotome, stained with 2% uranyl acetate for 2 h, and analysed with a transmission electron microscope (H-7500, Hitachi, Japan).

To measure the stomatal and pavement cell density and calculate the stomatal index, imprints were detached from the surface of the collected leaves and were mounted on a glass microscope slide (Nadakuduti et al., 2012). Samples were observed with an optical microscope (IX71, Olympus, Japan).

Water loss and chlorophyll leaching assay

Detached leaves of the gl6-ref mutant and wild-type were left at room temperature and photographed and weighed every 1 h. Water loss was calculated and represented by the percentage of fresh weight. Chlorophyll leaching assays were carried out as described (Aharoni et al., 2004). Briefly, excised seedling leaves were washed with tap water, weighed, and put in 30 ml of 80% ethanol at room temperature. Four hundred microliters was removed from each sample every 10 min and used to measure the absorbance at 664 and 647 nm wavelength. Finally, the samples were incubated in boiling water, cooled on ice, and used to examine the absorbance to evaluate the total chlorophyll content. The formula in the described method was used to calculate the chlorophyll content and chlorophyll extraction rate.

Drought tolerance experiment

Seeds of the BC₈F₃ homozygous gl6-ref mutant or wild-type were germinated in paper towels for 2 d and then transplanted in sand-filled pots in a greenhouse (27 °C day/23 °C night, 16 h day). The control plants were well watered, and the treatment plants were subjected to drought stress by withholding water for up to 20 d, and re-watered. Six biological replications were used for this assay, and the number of surviving seedlings used to calculate the survival rate. During the drought treatment, the relative water content and relative electrical conductivity were respectively monitored by a previously described method (Zheng et al., 2010). Control plants and experimental plants subjected to 10 d of drought stress were photographed. Thermal images were obtained using an infrared imaging system (VarioCam HD, InfraTec, Germany) following the manufacturer’s instructions.

Analysis of wax composition

Wax extraction and gas chromatography–mass spectrometry (GC-MS) analyses were performed according to the described methods with some modifications (Chen et al., 2011). The gl6 mutant and the wild-type were grown in a substrate of roseite and sand (1:1) in a growth chamber (25 °C) under 16/8 h light/dark for three leaves (about 6–7 d after planting). Five to six grams of fresh second leaves (6 d after planting) from gl6 mutant and wild-type seedlings were collected and immediately immersed in 5 ml chloroform for 45 s. The extracts containing 10 mg of tetracosane (Fluka) as an internal standard were transferred into opened reactive vials, dried with nitrogen gas (Pressure Blowing Concentrator; N-EVAP), derivatized by adding 20 μl of N,N-bis-trimethylsilyltrifluoroacetamide (Macherey-Nagel) and 20 μl of pyridine, and incubated for 40 min at 70°C. These derivatized samples were then analysed by GC with flame ionization detector (Agilent, Technologies) and GC-MS (Agilent gas chromatograph coupled to an Agilent 5973N quadrupole mass selective detector).

BSR-Seq, Seq-Walking and DLA analysis

About twenty glossy and non-glossy seedling segregants from the gl6-ref BC₈F₂ population were collected to construct mutant and non-mutant bulks for RNA extraction. Total RNA was isolated using the RNeasy Plus Mini Kit (Qiagen, USA) and RNA quality (RIN scores over 8) was checked on a Bioanalyzer 2100 (Agilent Technologies, USA) using an RNA 6000 Nano chip. RNA-Seq libraries were constructed using the Illumina RNA-Seq sample preparation kit according to the manufacturer’s protocol. The RNA-Seq libraries were sequenced on an Illumina HiSeq2000 instrument. Sequencing data were used to perform BSR-Seq analysis following our published method (Liu et al., 2012). For the Seq-Walking analysis, DNA was extracted using a cetyl trimethylammonium bromide method (Murray and Thompson, 1980) from the pools of heterozygous B73×(gl6-Mu/gl6-ref), in which half of the individuals contain the Mu-insertion allele, gl6-2 (Supplementary Table S1). Following quantification, DNA samples were sheared using a Biorupter machine (Diagenode, USA) and sequenced. The resulting sequence data were analysed according to our published Seq-Walking strategy (Li et al., 2013). DNA samples of gl6-2 were also used to perform DLA analysis according to our published method (Liu et al., 2009).

Transcriptome analysis

RNA-Seq data of the gl6 mutant and non-mutant bulks used in the BSR-Seq analysis were also used to compare transcriptome-wide differences between the mutant and its wild-type control. Reads were aligned to the maize reference genome B73 AGPv3 using Tophat2 (Kim et al., 2013). Transcript accumulation levels were calculated using the R package HTSeq (Anders et al., 2015) and differentially expressed genes were identified with the R package DESeq2 (Love et al., 2014). Tests for enrichment of differentially expressed genes were performed using agroGO with the standard settings (Fisher’s exact test with significance threshold of 0.05, Yekutieli multi-test adjustment) (Du et al., 2010).

qRT-PCR

For tissue-specific expression analysis, samples of young roots and leaves of 12 and 24 d after sowing (DAS), mature leaves of 12 and 27 d after pollination, husk, silk, anther, endosperm, embryo and seeds of 14 and 27 d after pollination were collected from the inbred line B73, and then were used to extract total RNAs by TRIzol reagent (Invitrogen). Quantitative real-time RT-PCR (qRT-PCR) analysis was conducted with TransStart Green qPCR SuperMix (TransGen Biotech) on a 7300-sequence detection system (Applied Biosystems). Maize actin1 was used as internal control, and the relative expression of gl6 mRNA was calculated using the 2^−ΔΔCtmethod (Livak and Schmittgen, 2001).

Subcellular location of GL6

To generate a GL6–yellow fluorescent protein (YFP) fusion construct, the open reading frame (ORF) of the Gl6 gene was cloned into the pEarleyGate-101 vector that contains the YFP reporter gene (Earley et al., 2006). Using previously described polyethylene glycol (PEG)-mediated transformation protocols (Chen et al., 2014), the GL6–YFP construct was transformed into maize protoplasts alone or co-transformed with an ER marker (red fluorescent protein (RFP)–CNX), a Golgi marker (mRFP–ManI), a trans-Golgi network (TGN) marker (RFP–SYP41), or a nuclear protein marker (mRFP–AHL22) (Xiao et al., 2009; Cui et al., 2014). Fluorescence signals were observed under a confocal laser scanning system (Leica Microsystems, Wetzlar, Germany).

Accession numbers

Sequence data from this article can be found in the GenBank/EMBL databases under the following accession numbers: GL6 (GRMZM2G139786). The accession numbers for proteins in the phylogenetic analysis are gi 670387534, gi 1162441486, gi 241943026, gi 514753272, gi 1238298764, gi 1070905880, gi 573963963, gi 125537042, gi 77556167, gi 222626102, gi 357161708, gi 1149739371, and gi 474448180.

Results

Morphological and biochemical characterization of the gl6 mutant

The spontaneous gl6 mutant first described by Emerson in 1935 has been designated gl6-ref (Emerson et al., 1935, Schnable et al., 1994). Like other glossy mutants in maize, seedling leaves of the gl6-ref mutant are shiny green in appearance, and water droplets easily form and adhere to leaf surfaces after leaves are sprayed with water (Fig. 1A). The accumulation of epicuticular waxes on the second leaf surface of the gl6-ref mutant was examined via FE-SEM. Substantially fewer wax crystals were observed on surfaces of gl6-ref mutant leaves relative to surfaces of wild-type leaves (Fig. 1B). Ultrastructural analysis of leaf epidermal cells by TEM found that gl6-ref epidermal cells contained irregular masses in the vacuole, which were absent from the wild-type cells (Fig. 1C, D). Within the masses, linear inclusions were found only in epidermal cells of the gl6-ref mutant (Fig. 1E). Other than this, no significant differences were observed between the two genotypes including cuticle morphology and the epidermal cell wall. This finding is similar to the observation of linear inclusions in Arabidopsis cer5 mutants, which exhibit defects in export of waxes through the PM (Pighin et al., 2004). Our results, therefore, suggested that waxes have accumulated within gl6 cells.

Fig. 1.

Morphological characterization of the gl6-ref mutant. (A) gl6-ref mutant exhibits a glossy phenotype. Water is sprayed on seedlings to distinguish gl6-ref mutant seedlings from wild-type (WT). (B) Adaxial leaf epicuticular wax accumulation in WT and mutant (gl6-ref) seedlings detected via SEM. ×5000 magnification. (C–E) TEM analysis of leaf epidermal wax secreting cells from WT and gl6-ref mutant. (E) Enlarged view showing the red rectangular area marked in (D). Arrows in (D, E) indicate unusual linear inclusions. Scale bars: 3 cm in (A), 5 µm in (B–D), 500 nm in (E). (This figure is available in color at JXB online.)

The wax load and composition of the gl6-ref mutant were assessed via GC-MS. Relative to wild-type, leaf surface wax loads on the gl6 mutant were reduced by 80%, from the wild-type level of 13.2 µg cm⁻² to 2.6 µg cm⁻² (Fig. 2A). Wax composition analysis showed that aldehydes and primary alcohols were highly reduced in the gl6 mutant, which decreased to 1.5% and 22% of wild-type levels, respectively. In addition, the accumulations of fatty acids, alkanes, ethyl stearate, β-sitosterol, and several unidentified wax classes were also decreased in the gl6 mutant (Fig. 2B). For individual wax constituents, amounts of C₃₂ aldehydes and C₃₂ primary alcohols on gl6 mutant leaves were significantly less than on wild-type leaves. Amounts of C₁₈, C₃₁ fatty acids, C₂₉, C₃₁, C₃₂ alkanes, C₃₀, C₃₄ aldehydes and C₃₀, C₃₃ primary alcohols were also reduced in the gl6 mutant (Fig. 2B). The compounds with reduced accumulation in the gl6 mutant are those having chain lengths of 29 or more carbons, with the exception of C₁₈ fatty acids. This suggests that gl6 has its main impact on the accumulation of VLCFAs and their derivatives.

Fig. 2.

Total leaf epicuticular wax load and wax composition of wild-type and gl6-ref mutant. (A) Total leaf cuticular wax load of wild-type and gl6-ref mutant. (B) Epicuticular wax composition of wild-type and gl6-ref mutant. Values are means of eight biological replicates ±SD. UI, unidentified. Asterisks indicate statistically significant differences between wild-type and gl6-ref mutant (*P<0.05, **P<0.01, ***P<0.001, Student’s t-test).

We further quantified total waxes including both surface and intracellular waxes and determined the proportion of total waxes secreted to the surface. This experiment was performed according to published methods (McFarlane et al., 2014). Briefly, two separate approaches were used to collect (i) leaf surface waxes, representing epicuticular waxes, and (ii) leaf total waxes, including both surface/epicuticular and intracellular waxes. Surface waxes were extracted through brief dipping of plant leaves in chloroform. In contrast total waxes were extracted by homogenizing leaves in chloroform. Extracted waxes were quantified via GC-MS. As a result, total waxes were reduced from the wild-type level of 19.7 µg cm⁻² to 13.1 µg cm⁻² in the gl6 mutant (Table 1). In wild-type leaves, 67.1% of total waxes were from surface waxes, suggesting that most waxes were secreted (Table 1). In contrast, only 19.9% of total waxes were surface waxes in mutant leaves. In conjunction with the observation of linear inclusions in gl6 epidermal cells (Fig. 1D, E), these results suggest that the wax exporting system is dysfunctional, causing a large proportion of waxes to remain within epidermal cells. Taken together, these results indicated that gl6 is involved in both the biosynthesis of very-long-chain waxes, particularly C₃₂ aldehydes and C₃₂ primary alcohols, and, possibly, wax transport from the ER to the plant surface.

Table 1.

Epicuticular wax extracted from the leaf surface compared with total wax of wild-type and gl6 mutant

Parameter	WT	gl6-ref
Total wax (µg cm−2)	19.7±4.1	13.1±2.4
Surface wax (µg cm−2)	13.2±2.3	2.6±0.5
Wax secretion ratio	67.1%	19.9%

Total wax extraction and calculation of ratio of wax secretion were performed following a reported measurement strategy (McFarlane et al., 2014). Total and surface wax content are means ±SD.

Increased cuticle permeability and drought sensitivity of the gl6 mutant

Reduced cuticular wax accumulation is associated with increased cuticle permeability (Kerstiens, 1996; Aharoni et al., 2004; Bessire et al., 2007). We performed chlorophyll leaching and water loss assays using seedling leaves of gl6-ref mutant and wild-type plants. For the chlorophyll leaching assay, the concentration of leaf chlorophyll in the solution was monitored at various time points. The results showed that leaf chlorophyll leaching of the gl6-ref mutant was faster than that of wild-type (Fig. 3A), even though the total leaf chlorophyll contents of the gl6-ref mutant and wild-type were similar (Fig. 3B). To detect water loss, detached leaves of gl6-ref and wild-type were exposed to air at room temperature; gl6-ref mutant leaves were obviously curled and wilted after only 2 h (Fig. 3C). The rate of water loss of detached leaves from the gl6-ref mutant was significantly higher than that of wild-type (Fig. 3D). Results from both the chlorophyll leaching and water loss assays suggested that gl6-ref mutant leaves exhibit increased cuticle permeability as compared with wild-type leaves.

Fig. 3.

Leaf surface permeability analysis of gl6-ref mutant. (A) Kinetics of chlorophyll leaching from leaves of the WT and gl6-ref mutant. (B) The total leaf chlorophyll content of WT and gl6-ref mutant. (C) Detached leaves from WT and gl6-ref mutant immediately after harvest and after 2 h at room temperature. (D) Water loss of detached leaves of the WT and gl6-ref mutant. Values in (A, B, D) are means of three biological replicates ±SD. (This figure is available in color at JXB online.)

Leaf water loss has been associated with reduced leaf surface temperature, and hence monitoring leaf surface temperature is widely used as an indicator of leaf water loss (Mustilli et al., 2002). The surface temperature of gl6-ref mutant leaves, monitored via infrared thermography, was lower than that of wild-type leaves under both drought stress and well-watered conditions (Fig. 4A), while stomatal density, pavement cell density, and stomatal index of the gl6-ref mutant were similar to those of wild-type (Fig. 4B–D). These results suggested that the decreased leaf wax accumulation of the gl6-ref mutant caused faster water losses.

Fig. 4.

gl6-ref mutant seedlings are sensitive to drought stress. (A) False-color infrared image of the wild-type and the gl6-ref mutant under well-watered and drought stressed conditions. (B–D) Stomatal density (B), pavement cell density (C), and stomatal index (number of stomata per total epidermal cells) (D) analysed in leaf abaxial epidermal layers from WT and gl6-ref mutant. Data are means of five individual plants. (E) Drought phenotypes of WT and gl6-ref mutant seedlings in soil following drought stress and after re-watering. (F) The survival rate of WT and gl6-ref mutant seedlings after drought stress and re-watering. (G, H) The leaf relative water content and relative electrical conductivity of WT and gl6-ref mutant seedlings under well-watered (WW), drought stressed (DS) and re-watered (RW) conditions. In (B–D, F), data are means of five replicates ±SD, and in (G, H) data are means of three replicates ±SD (***P<0.001, Student’s t-test). Scale bars: 10 cm in (A, E). (This figure is available in color at JXB online.)

Given the increased cuticular permeability and leaf water loss of the gl6-ref mutant, we investigated how gl6-ref mutant seedlings respond to drought stress. As a result, under severe drought stress, the gl6-ref mutant showed a more severe wilting phenotype as compared with wild-type controls (Fig. 4E), and after re-watering, about 50% of wild-type seedlings could recover and survive, whereas none of the gl6-ref mutant seedling plants could survive (Fig. 4E, F). Under the well-watered conditions, no significant phenotypic differences in either the wilting phenotype or the relative water content of leaves were observed between gl6-ref mutant and wild-type seedlings (Fig. 4E, G). In contrast, under mild drought stress, the relative water content of gl6-ref mutant seedling leaves was lower than that of wild-type controls, 53.0% in gl6-ref as compared with 72.3% in wild-type. The re-watering plants after drought stress showed no significant differences in the leaf relative water content between two genotypes (Fig. 4G). In addition, a leaf electrolyte leakage assay for evaluating membrane damage was conducted, and the results showed that, under mild drought stress, the gl6-ref mutant exhibited an electrical conductivity of 35.1%, higher than the value of 18.2% of wild-type (Fig. 4H), which indicated that a larger degree of membrane damage in the mutant resulted in higher electrolyte leakage. In re-watered plants, no significant differences of the relative conductivity were observed between mutant and wild-type (Fig. 4H). The results corroborated that the gl6-ref mutation reduced seedling tolerance to drought.

Molecular cloning of gl6

Previously, the gl6 gene had been mapped roughly to the long arm of chromosome 3 (Schnable et al., 1994). BSR-Seq, a method utilizing RNA-Seq for bulked segregant analysis (Liu et al., 2012), was performed to map the gene. Briefly, segregating BC₈F₂ populations derived from self-pollination of gl6-ref heterozygous plants were used to identify and collect mutant and wild-type plants, separately pooled into two bulks for RNA extraction and sequencing. Three replicates were conducted, producing six RNA-Seq datasets, each of which had more than 20 million reads. RNA-Seq reads were aligned to the B73 reference genome (AGPv2), and the polymorphic single nucleotide polymorphisms identified were used as genetic markers to map the target gene. Through this process, the gl6 gene was mapped to an 18.9 Mb interval on chromosome 3, from 113.5 to 132.4 Mb (Fig. 5A).

Fig. 5.

Molecular cloning of the gl6 gene and its phylogenetic analysis. (A) BSR-Seq analysis of an F₂ segregated gl6-ref population mapped gl6 to the 113.5–132.4 Mb interval of chr3. (B) PAGE results of digestion–ligation–amplification (DLA) analysis using the adaptor primer (Nsp-15ctc). The rectangle indicates the specific bands produced from gl6/+ plants but not from WT plants. (C) The gene structure of gl6, Mu insertions in four alleles, and lesions in two gl6 EMS alleles. (D) Phylogenetic tree constructed using MEGA 7.0 and the GL6 protein and GL6 homologs aligned using ClustalW. Distances were estimated with a neighbor-joining algorithm, and bootstrap support is indicated to the left of branches. (This figure is available in color at JXB online.)

To facilitate the identification of the gl6 gene, additional gl6 alleles were generated through Mu transposon mutagenesis and chemical (EMS) treatment (Candela and Hake, 2008). Four new Mu-tagged alleles were identified in the progeny of crosses between homozygous gl6-ref mutant and Mu-active lines; these mutants were designated gl6-2, gl6-3, gl6-4, and gl6-5 (see Supplementary Table S1). An EMS mutagenesis screen identified three glossy mutants that were verified to be allelic to gl6-ref, and which were termed gl6-6, gl6-7, and gl6-8 (Supplementary Table S1).

One Mu-tagged allele, gl6-2, was used to perform Seq-Walking sequencing, a genome walking approach to amplify and isolate DNA fragments flanking Mu transposon insertions throughout the genome (Li et al., 2013). In total, 28 non-redundant Mu flanking sequence (MFS) sites with >30 reads were identified in the 18.9 Mb BSR-Seq mapping interval, and one Mu-insertion hotspot gene, GRMZM2G139786, containing three MFSs was identified (see Supplementary Fig. S1; Supplementary Table S2). At the same time, the gl6-2 allele was also subjected to DLA analysis, an adaptor-mediated PCR-based method for isolating Mu flanking sequences that co-segregated with mutant phenotypes (Liu et al., 2009). The DLA result independently identified a Mu insertion in GRMZM2G139786 that co-segregated with the gl6 glossy phenotype (Fig. 5B). Based on these results, all four Mu-derived alleles, gl6-2, gl6-3, gl6-4, and gl6-5, were sequenced to identify mutations in GRMZM2G139786. Mu1 insertions were identified 37 bp upstream of the gl6 start codon in the gl6-2 allele, 211 bp upstream of the start codon in both gl6-4 and gl6-5 alleles (Dietrich et al., 2002), and 320 bp downstream of the start codon in the gl6-3 allele (Fig. 5C).

The genomic sequences of GRMZM2G139786 from three EMS-induced gl6 alleles were amplified and Sanger sequenced (see Supplementary Table S1). These analyses showed that the gl6-6 allele contained a G to A transition 314 bp downstream of the start codon, producing a premature termination codon (PTC); gl6-7 carried a C to T transition 391 bp downstream of the start codon, causing an amino acid change from arginine to cysteine; and the gl6-8 allele contained a 9-bp deletion 497 bp downstream of the start codon and seven non-synonymous mutations in the coding region (Supplementary Table S1; Fig. 5C). In addition, based on the BSR-Seq result, the accumulation of the GRMZM2G139786 transcripts was significantly reduced in the gl6-ref mutant pool as compared with the wild-type pool (Supplementary Table S3). Collectively, these results demonstrate that GRMZM2G139786 is the gl6 gene.

Characterization of the novel glossy gene, gl6

The gl6 gene has a single exon with a 675-bp open reading frame and encodes a putative protein with 224 amino acids. Domain analysis showed that GL6 is a novel protein containing a conserved DUF538 (domain of unknown function) between positions 29 and 138 aa of the protein (see Supplementary Fig. S2). Homologs of GL6 can be identified in sorghum, rice, Arabidopsis, and other plants. Phylogenetic analysis using the full-length protein sequences of GL6 homologs showed that GL6 was grouped into a monocot-specific subfamily (Fig. 5D), and multiple sequence alignment showed that the DUF538 domain of these proteins is highly conserved (Supplementary Fig. S2).

A fusion construct of GL6 with C-terminal YFP was generated and expressed in maize protoplast cells. Observation via confocal microscopy showed that GL6–YFP fluorescence signals were detected in the cytoplasm and the plasma membrane, but not in the nucleus, whereas the control YFP signal was observed throughout the cell (Fig. 6A, B). To further study the compartmental localization of the GL6 protein, we co-expressed GL6–YFP with the ER marker red fluorescent protein (RFP)–CNX, the Golgi marker mRFP–ManI, the trans-Golgi network (TGN) marker RFP–SYP41 and the nuclear protein marker mRFP–AHL22 in the protoplast cells, and the results revealed that GL6–YFP co-localized with the ER, Golgi, and TGN markers (Fig. 6C–E), but not with the nuclear protein marker (see Supplementary Fig. S3). To confirm plasma membrane localization of GL6, the GL6–YFP-expressing cells were incubated with the plasma membrane marker FM4-64, and the result showed that the signals of GL6–YFP and FM4-64 co-localized as expected (Fig. 6F). Together, our evidence indicated that GL6 is an ER membrane-, Golgi-, TGN-, and plasma membrane-localized protein.

Fig. 6.

Subcellular localization of the GL6 protein in maize protoplasts. Confocal images show the expression of YFP, GL6 fused at its C terminus with YFP, and organelle markers. (A, B) Subcellular localization YFP and GL6–YFP. (C) Co-localization of GL6–YFP with RFP–CNX (an ER marker). (D) Co-localization of GL6–YFP with mRFP–ManI (a Golgi marker). (E) Co-localization of GL6–YFP with RFP–SYP41 (a TGN marker). (F) Co-localization of GL6–YFP with FM4-64 (a plasma membrane marker). Scale bar: 10 µm. (This figure is available in color at JXB online.)

gl6 expression and transcriptome analysis of the gl6 mutant

Real-time quantitative RT-PCR analysis was performed to detect gl6 gene expression in different maize tissues, and the results showed that gl6 was significantly expressed in leaves and silk, especially in young leaf, but there was lower expression in root, husk, anther, and immature seed (see Supplementary Fig. S4). To further understand the impact of the gl6 mutation on the transcriptome, we analysed differential expression in the gl6 mutant and wild-type using the BSR-Seq data. With a 10% false discovery rate, 421 differentially expressed (DE) genes were identified (Supplementary Table S4). Among the DE genes, 235 and 186 genes were up- and down-regulated in the gl6 mutant versus wild-type, respectively (Supplementary Fig. S5A). Among the 235 up-regulated or 186 down-regulated genes, 197 (84%) and 135 (73%) genes exhibited greater than 2-fold changes. Gene Ontology (GO) enrichment analysis was performed on these DE genes using AgriGO (Du et al., 2010), and complete lists of significantly enriched GO terms are shown in Supplementary Table S4. Significant enrichment of cellular response to stimulus and stress, especially to water and heat stresses, were observed among the enriched GO terms (Supplementary Fig. S5B), which is consistent with the hypothesis that the reduced wax accumulation on gl6 leaves results in increased sensitivity to stimuli and stresses. Moreover, an enrichment of GO terms related to the fatty acid biosynthetic process was also observed. Of genes in these GO terms, two genes, GRMZM2G029912 and GRMZM2G083526, which are homologs of the Arabidopsis CER3/WAX2 gene (AT5G57800) involved in the biosynthesis of VLCFAs, and another gene, GRMZM2G031790, a homolog of the Arabidopsis KCS2 gene (AT1G04220), showed significantly higher expression in the gl6 mutant with 12.0-, 4.8- and 12.5-fold changes, respectively (Lee et al., 2009; Bernard et al., 2012; Supplementary Table S3). This result suggested that altering wax amount and localization in the gl6 mutant resulted in feedback, causing up-regulation of some genes in the fatty acid biosynthesis pathway.

Discussion

In this study, multiple strategies were combined to facilitate the genetic mapping and identification of a gl6 candidate gene, which was confirmed using multiple independent Mu-tagged and EMS-induced gl6 alleles. gl6 is a novel gene with a conserved uncharacterized domain, opening the door to extending our understanding of the molecular basis of cuticular wax accumulation.

The phenotype and physiological characterization of the gl6 mutant showed that gl6 is a typical maize glossy mutant with reduced epicuticular wax accumulation on the leaf surface (Figs 1, 3). The amount of total cutin monomer present in gl6-ref mutant and wild-type seedling leaves was also examined, but no significant differences were detected, which indicates that the role of the gl6 gene is specific to epicuticular waxes. Cuticular waxes, the hydrophobic layer on plant surfaces, protect leaves from non-stomatal water loss (Samuels et al., 2008). However, few studies provide quantitative data to measure drought tolerance of maize glossy mutants. We demonstrated that the reduction of epicuticular waxes in the gl6 mutant was associated with increased cuticle permeability and reduced drought tolerance of maize seedlings (Figs 3A, 4E). Studies from other plant species have shown that increases in cuticular wax load can enhance drought tolerance (Lü et al., 2012; Luo et al., 2013; Zhou et al., 2013; Zhu and Xiong, 2013; Al-Abdallat et al., 2014). Future studies should test the impact of elevated expression of cuticular wax on plant drought tolerance in adult maize plants as well as the potential physiological cost of accumulating abundant cuticular waxes.

Sequence analysis revealed that gl6 encodes a novel protein containing a conserved DUF538 domain with no defined function (Supplementary Fig. S2). gl6 orthologs were identified in sorghum, rice, Arabidopsis, and other plants, but none has been characterized. However, plant DUF538 proteins are predicted to be the potential homolog of mammalian BPI (bactericidal/permeability increasing) proteins (Gholizadeh and Kohnehrouz, 2013), which have been suggested to be important for the defense system against various pathogens (Srivastava et al., 2007). Furthermore, BPI proteins are predicted to be structurally similar to esterase-type hydrolases, which implies DUF538 might have a lipid-associated enzymatic activity (Gholizadeh, 2014, 2016). The potential role of the gl6 gene in wax transportation may result from the interaction of the GL6 protein with other transporters, as has been shown in other systems (Park et al., 2013). In this study, we determined that maize gl6 is involved in wax accumulation and drought tolerance, which will provide insights into understanding the molecular function of DUF538 family members in plants. It has been reported that the maize gl1 mutant shows smaller and more densely arrayed trichomes (Sturaro et al., 2005). Mutants of the Arabidopsis gl1 homolog (yre-1) exhibit a wax-deficient phenotype that is associated with organ fusion and various abnormalities in trichome development (Kurata et al., 2003). A T-DNA insertion mutation of the Arabidopsis gl6 homolog, At1g56580, svb-1, exhibits smaller trichomes with variable branches as compared with wild-type (Marks et al., 2009). However, no phenotype associated with cuticular wax accumulation has been reported for the svb-1 mutant.

The gl6 gene encodes a novel protein. Our characterizations have provided clues as to its function. VLCFAs are synthesized in the ER but the processes by which wax components from the ER are delivered to the plasma membrane remain unknown. Two hypotheses have been proposed, a Golgi-mediated vesicular trafficking pathway and transfers via physical ER–PM connections (Kunst and Samuels, 2003; Schulz and Frommer, 2004). Two vesicle-trafficking genes, GNL1 and ECH, have been implicated in wax transport from the ER to the PM (McFarlane et al., 2014), which supports the Golgi-mediated vesicular trafficking hypothesis. In the current study, the GL6 protein was localized in the ER membrane, the Golgi, the TGN, and the plasma membrane (Fig. 6), which provides further support for the involvement of the Golgi in intracellular wax transport. Our results also suggested that GL6 might play a role in the process of extracellular wax transport. Quantification of secreted waxes on the leaf surface and total leaf waxes showed that a decrease of total waxes in the mutant was largely attributable to reduced accumulation of surface waxes (Table 1). Ignoring surface waxes, the remaining wax accumulation in mutants was higher than that in wild-type, indicative of dysfunctional wax export to leaf surfaces in the gl6 mutant. Linear inclusions found in epidermal cells of the gl6 mutant via TEM analysis provided further evidence that waxes accumulate inside epidermal cells of the gl6 mutant (Fig. 1E). Taken together, our results suggest that GL6 functions in the intracellular transport of cuticular waxes, thereby influencing cuticular wax accumulation.

Supplementary data

Supplementary data are available at JXB online.

Fig. S1. Mu insertion sites from Seq-Walking.

Fig. S2. Multiple sequence alignment of amino acid sequences of the predicted maize GL6 protein and its homologs in other species.

Fig. S3. Co-localization of GL6–YFP with mRFP–AHL22 (nuclear marker).

Fig. S4. Accumulation of gl6 transcripts in various tissues.

Fig. S5. Summary of DE genes and GO analysis.

Table S1. gl6 allele information and the corresponding mutation types.

Table S2. Mu flanking sequences and Mu sequences for each gl6 Mu tagging allele.

Table S3. Differentially expressed genes in gl6 mutant versus wild-type.

Table S4. Enriched GO terms in gl6 mutant differentially expressed genes.