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Plant Cell Physiol
Plant Cell Physiol
pcp
Plant and Cell Physiology
0032-0781
1471-9053
Oxford University Press UK

38757823
10.1093/pcp/pcae053
pcae053
Regular Paper
AcademicSubjects/SCI01180
Micromorphological and Chemical Characterization of Drimys winteri Leaf Surfaces: The Secondary Alcohols Forming Epicuticular Wax Crystals Are Accompanied by Alkanediol, Alkanetriol and Ketol Derivatives
Zhang Zhonghang Department of Botany, University of British Columbia, 6270 University Boulevard, Vancouver, BC V6T 1Z4, Canada

Mistry Dwiti Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, BC V6T 1Z1, Canada

https://orcid.org/0000-0002-6302-2835
Jetter Reinhard Department of Botany, University of British Columbia, 6270 University Boulevard, Vancouver, BC V6T 1Z4, Canada
Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, BC V6T 1Z1, Canada

*Corresponding author: E-mail, reinhard.jetter@ubc.ca
8 2024
17 5 2024
17 5 2024
65 8 12451260
08 9 2023
22 4 2024
14 5 2024
13 5 2024
13 6 2024
© The Author(s) 2024. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
2024
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Abstract

The cuticle is a hydrophobic coating of most aerial plant surfaces crucial for limiting non-stomatal water loss. Plant cuticles consist of the lipid polyester cutin and associated waxes with compositions varying widely between plant species and organs. Here, we aimed to provide a comparative analysis of the dark-glossy adaxial and pale-glaucous abaxial sides of Drimys winteri (Winteraceae) leaves. Scanning electron microscopy showed nanotubular wax crystals lining the entire abaxial side of the leaf (including stomatal pores), while the adaxial side had patches of mixed platelet/tubule crystals and smooth areas between them. Consecutive treatments for wax removal and cutin depolymerization revealed that the waxes were deposited on a cutin network with micron-scale cavities across the entire abaxial surface including the stomata pores, and on a microscopically smooth cutin surface on the adaxial side of the leaf. Gas chromatography coupled to mass spectrometry and flame ionization detection showed that the wax mixtures on both sides of the leaf were complex mixtures of very-long-chain compounds dominated by the secondary alcohol nonacosan-10-ol and alkanediols with one hydroxyl on C-10. It is therefore very likely that the characteristic tubular wax crystals of both leaf sides are formed by these alcohols and diols. Further secondary alcohols and alkanediols, as well as ketols and alkanetriols with one functional group on C-10, were identified based on mass spectral fragmentation patterns. The similarities between all these mid-chain-functionalized compounds suggest that they are derived from nonacosan-10-ol via regio-specific hydroxylation reactions, likely catalyzed by three P450-dependent monooxygenases with different regio-specificities.

Cuticular wax
Cutin
Mass spectrometry
Nonacosan-10-ol
Scanning electron microscopy
Stomatal plugs
Natural Science and Engineering Council (Canada) Discovery Grants Program 262461 Natural Science and Engineering Council (Canada) Discovery Grants Program 262461
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pmcIntroduction

The aerial surfaces of land plants are covered by a hydrophobic coating produced by their epidermal cells. This cuticle plays important roles in plant development and adaptation to stresses, especially in limiting non-stomatal water loss (Yeats and Rose 2013). The core framework of the cuticle is established by cutin, an ester-linked polymer of hydroxylated and epoxidated C16 and C18 fatty acids and glycerol (Fich et al. 2016). It is associated with cuticular waxes, which are either deposited within the polymer as intracuticular wax or onto its outer surface as epicuticular wax (Buschhaus and Jetter 2011). Cuticular waxes are mixtures of very-long-chain (VLC, C20 to C40) derivatives of fatty acids, frequently also comprising cyclic compounds such as triterpenoids and phenolics (Jetter et al. 2006). The qualitative and quantitative compositions of cuticular wax mixtures vary widely between plant species and organs.

The biosynthesis of common cuticular wax compounds has been investigated in detail, mainly in the model plant Arabidopsis thaliana. The wax biosynthetic pathways reside in the ER of epidermal cells, where they convert fatty acids supplied by plastids by first extending their hydrocarbon backbones and then modifying the acyl head groups (Samuels et al. 2008). In the process, C16 acyl-CoA substrate has its hydrocarbon tail first extended by fatty acid elongase complexes consisting of four independent enzymes (Samuels et al. 2008): (i) a 3-ketoacyl-CoA synthase (KCS) condensing the acyl-CoA starter with a two-carbon unit from malonyl-CoA into a 3-ketoacyl-CoA (Joubès et al. 2008), (ii) a 3-ketoacyl-CoA reductase (KCR) forming the corresponding 3-hydroxyacyl-CoA (Beaudoin et al. 2009), (iii) a 3-hydroxyacyl-CoA dehydratase forming a trans-2,3-enoyl-CoA (Bach et al. 2008) and (iv) an enoyl-CoA reductase yielding an acyl-CoA two carbons longer than the starter (Zheng et al. 2005). This reaction cycle is repeated until the acyl reaches chain lengths up to C40, with a characteristic chain length distribution typically peaking around C30. Respective VLC acyl-CoAs then may have their head groups modified by a fatty acyl-CoA reductase into corresponding primary alcohols (Rowland et al. 2006), and the latter can be combined with fatty acyl-CoAs to form wax esters (Li et al. 2008). Alternatively, the VLC fatty acyl-CoAs can also be converted into aldehydes and then decarbonylated into alkanes by the CER1 and CER3 enzymes (Aarts et al. 1995, Bernard et al. 2012). The alkanes may finally be hydroxylated by a P450-dependent enzyme, mid-chain alkane hydroxylase (MAH1), into corresponding secondary alcohols, diols, ketones and ketols with functional groups residing mainly on C-15 and C-14 (Greer et al. 2007).

The common wax compounds are thought to make up the bulk of the intracuticular wax in many species (Buschhaus and Jetter 2012) and thus contribute largely to the main cuticle function of limiting transpirational water loss (Buschhaus and Jetter 2012, Jetter and Riederer 2016). More of the same compounds then typically form a smooth film across the epidermal cell surfaces, giving the overall organs a glossy appearance (Traore et al. 1989). However, if one or several wax components accumulate to high concentration, they may form microscopic crystals that scatter light and make the surface glaucous (Peterson et al. 1982). Scanning electron microscopy (SEM) showed that these epicuticular wax crystals have characteristic shapes and arrangements largely determined by their composition (Barthlott et al. 1998). For example, the primary alcohols dominating the surface wax of wheat form platelet-shaped crystals (Koch et al. 2006), while wax aldehydes form platelet crystals on rice leaves (Haas et al. 2001).

Wax crystals with characteristic nanotubule shapes were reported for species in diverse families, including mosses (Robinson 1971), ferns (Guo et al. 2018), gymnosperms (Pinaceae and Taxaceae) (Franich et al. 1978, Wen et al. 2006) and early-diverging angiosperms (Papaveraceae, Nelumbonaceae and Tropaeolaceae) (Jetter and Riederer 1996, Koch et al. 2006). The occurrence of respective nanotubules was tightly linked to the accumulation of nonacosan-10-ol (Barthlott et al. 1998), suggesting that these crystals are formed by this unique secondary alcohol. This conclusion was corroborated by experiments, showing that nanotubles removed mechanically from the plant surface consist mainly of nonacosan-10-ol (Holloway et al. 1976) and that nonacosan-10-ol crystallizes in vitro into nanotubules independent of biological factors (Jeffree et al. 1975, Jetter and Riederer 1994). Nonacosan-10-ol is typically associated with structurally similar alkanediols that can also form nanotubules and are likely mixed into the crystals found on respective plant surfaces (Jetter and Riederer 1995).

The presence of epicuticular wax crystals greatly affects the properties of the plant surface, and the crystals are thought to serve ecophysiological functions including protection from insect herbivores, pathogen spores and dust (Ensikat et al. 2011, Yeats and Rose 2013). In addition, the three-dimensional networks of nanotubules filling the cavity over sunken stomata are thought to play special roles associated with ecophysiological adaptations, most prominently of gymnosperm needles (Jeffree et al. 1971, Brodribb and Hill 1997, Šantrůček 2022). The functions of these so-called stomatal plugs rely on their porous structure, allowing for gas exchange while preventing liquid penetration or accumulation. On the one hand, the stomatal plugs found on many conifer needles are thought to contribute to drought resistance, both by reducing residual transpiration from closed stomata (Šantrůček 2022) and by limiting water loss from open stomata (Jeffree et al. 1971). On the other hand, the stomatal wax plugs can play a role in keeping stomata dry, where they thus maintain free gas exchange in cold, humid environments (Brodribb and Hill 1997) and also impede growth of fungal hyphae toward and into the pores (Mohammadian et al. 2009). It is well established that the stomatal plugs on gymnosperm needles are formed by nanotubules consisting of nonacosan-10-ol (Jetter and Riederer 1994, Jeffree 2006, Wen et al. 2006).

Special stomatal architectures, including sunken stomata and epistomatal plugs, have also been reported for the abaxial leaf surface of Drimys winteri (Winteraceae) (Bongers 1973, Feild et al. 1998), a small angiosperm tree growing in the cloud forest of the southern Andes (Smith 1945). In contrast to the stomatal plugs of gymnosperm needles, the function of D. winteri is thought to be mainly in keeping the leaf surface above the pores dry, and not in moderating gas exchange for drought resistance (Feild et al. 1998). In this context, it is of interest that the stomatal plugs of D. winteri leaves were described as cutinaceous rather than waxy material (Bailey and Nast 1944, Bongers 1973).

Despite the functional importance of wax constituents like nonacosan-10-ol, the biosynthetic mechanisms forming them have not been investigated. It thus remains unclear whether the metabolic pathways leading to the Arabidopsis wax secondary alcohols (nonacosan-14- and 15-ol) may also form different alcohol structures in species of other families. Detailed information on the secondary alcohols accumulating in the wax mixtures of diverse plant taxa may help narrow down possible biosynthesis mechanisms. Therefore, the goal of the present study was to provide a comprehensive analysis of the cuticular waxes from a family lacking prior information on wax composition. We focused on D. winteri, since previous SEM investigations had documented the presence of epicuticular wax nanotubules typically associated with nonacosan-10-ol, at least on part of its leaf surface (Barthlott and Theisen 1994). The stark contrast between the dark-glossy adaxial and pale-glaucous abaxial sides of the D. winteri leaves (Supplementary Fig. S1) (Carson 1847) enabled comparative investigations between two surfaces of the same organ. In particular, we aimed to analyze the cuticle micromorphology on both sides of the leaf using light microscopy and SEM, and the chemical composition of the wax mixtures lining them using gas chromatography with mass spectrometry (GC–MS) and flame ionization detector (GC–FID).

Results

This investigation aimed to characterize the waxes lining the adaxial and abaxial leaf surfaces of D. winteri by (i) observing their micromorphology with light microscopy and SEM, (ii) analyzing the overall wax mixtures extracted from them by GC–MS and (iii) quantifying the detailed chain length and isomer profiles of individual wax fractions.

Micromorphology of the adaxial and abaxial leaf surfaces of D. winteri

To characterize the micromorphology of epicuticular waxes covering leaves of D. winteri, both sides of mature leaves were investigated using SEM. The adaxial leaf surface was relatively uniform and level overall, structured only with broad ridges that likely reflected anticlinal walls of underlying epidermis cells (Fig. 1A). Both the ridges and the areas between them comprised patches of epicuticular wax crystals and small smooth areas between them (Fig. 1B, C). The crystals mostly lacked sharp edges and distinct shapes, but some of them had hollow openings suggesting tubules 1–3 µm long and 0.1–0.2 µm wide (Fig. 1C). The abaxial leaf surface was also smooth overall, with shallow humps approximately 30 µm in diameter (Fig. 1D, E). Both the humps and the areas between them were covered by a dense network of epicuticular wax crystals (Fig. 1D). The crystals on areas outside the humps were slightly curved tubules 0.5–2 µm long and 0.1–0.4 µm wide, with relatively big air spaces between them (Fig. 1F, G). The crystals forming the humps were also tubules, with diameters similar to those outside the humps but only 0.5–1 µm long (Fig. 1H, I). They were frequently fused into clumps and had smaller air spaces in between. Overall, both sides of the D. winteri leaf thus had clearly different epicuticular wax crystal micro-morphologies, underscoring respective differences in the macroscopic appearance of the two surfaces.

Fig. 1 Micromorphology of intact D. winteri leaf surfaces. Scanning electron micrographs of (A–C) the adaxial side of the leaf and (D–I) the abaxial leaf surface. (A, D, E) Overall appearance of the surfaces and (B, C, F–I) high-magnification views of the epicuticular wax crystals. (F, G) Wax crystals in areas outside the surface humps. (H, I) Wax crystals within the surface humps. Images shown are representative, and two repeats gave similar results. Bars = 200 μm (E), 50 μm (A, D), 20 μm (B, F, H) and 2 μm (C, G, I).

The humps on the abaxial surface of D. winteri leaves suggested stomatal wax plugs, implying that different epidermal cell types may have epicuticular wax crystals with distinct shapes or arrangements. To test this, we used adhesive tape to peel exposed surface lipids and visualized the underlying surface with SEM. On the abaxial side of the leaf, the adhesive preferentially removed the protruding circular humps, revealing stomata underneath (Supplementary Fig. S2A, B) and confirming that the humps were plugs of extracellular material deposited over the stomata. Similar adhesive treatment of the adaxial surface did not remove sufficient material to expose the underlying cell surface.

To further test the composition of the stomatal plugs, we investigated the micro-morphology of D. winteri leaf surfaces after treatments designed to dissolve the surface waxes or the cutin polymer accompanying them. After rinsing the leaf with chloroform, a solvent known to remove waxes (Riederer and Schneider 1989), the adaxial surface still showed ridges outlining the underlying epidermal cells (Fig. 2A). However, the de-waxed surfaces of both the ridges and the areas between them had an irregular sub-micron relief instead of the wax crystals (Fig. 2B, C). Further treatment with methanolic sulfuric acid, known to dissolve the polyester cutin (Bonaventure et al. 2004), left the ridges unchanged (Fig. 2D) but smoothened the micro-relief on the entire adaxial surface (Fig. 2E, F). Removal of the wax from the abaxial side of the leaf did not affect the smooth overall appearance of the surface (Fig. 2G) but revealed a reticulate microscopic texture (Fig. 2H). The network structure was relatively dense in the circular stomatal plugs (Fig. 2H), while showing sub-micron cavities mostly in areas surrounding them (Fig. 2I). Consecutive cutin removal left ridges (Fig. 2J) and smooth surfaces similar to those on the adaxial side. However, the likely plug areas now had small, irregular ledges protruding from a smooth surface (Fig. 2K, L). Neither the solvent rinses nor the cutin depolymerization treatments exposed the stomata previously detected on the abaxial leaf surface.

Fig. 2 Micromorphology of D. winteri leaf surfaces treated to remove wax and/or cutin. Scanning electron micrographs of (A–F) the adaxial side of the leaf and (G–L) the abaxial leaf surface. Leaf surfaces after (A–C, G–I) extraction of waxes with chloroform and (D–F, J–L) further treatment with reagents removing cutin. Images shown are representative, and two repeats gave similar results. Bars = 50 μm (A, D, G, J), 20 μm (B, E, H, K) and 2 μm (C, F, I, L).

To assess stomatal densities, both sides of the D. winteri leaf were investigated using light microscopy. On the adaxial side of the leaf, pavement cells were clearly distinguishable and stomata could not be found (Supplementary Fig. S3A–C). In contrast, stomata were visible on the abaxial surface, along with clearly delineated pavement cells (Supplementary Fig. S3D–F). Solvent rinses to remove the waxes did not change epidermal cell geometries but facilitated stomata counts (Supplementary Fig. S4). The stomata density on the abaxial side of the leaf was 179 ± 2.4 mm−2.

Composition of overall cuticular wax mixtures covering the adaxial and abaxial leaf surfaces of D. winteri

To assess the chemistry underlying the different wax mircoreliefs on both sides of the D. winteri leaf, cuticular waxes were extracted separately from the adaxial and abaxial surfaces for chemical analysis. Wax compounds were identified by GC–MS and quantified by GC–FID (using a known amount of internal standard). The abaxial leaf side was covered by 31.3 ± 0.9 µg cm−2 of wax, of which 1.8 ± 0.4 µg cm−2 remained unidentified (Fig. 3). The abaxial wax mixture consisted predominantly of secondary alcohols (10.7 ± 0.1 µg cm−2) and alkanediols (12.6 ± 0.5 µg cm−2), along with primary alcohols (4.0 ± 0.2 µg cm−2) and p-coumarates (2.2 ± 1.0 µg cm−2). In contrast, the adaxial side of the leaf had a wax coverage of 9.0 ± 0.4, comprising approximately equal amounts of secondary alcohols (2.0 ± 0.1 µg cm−2), primary alcohols (2.2 ± 0.1 µg cm−2), alkyl esters (2.0 ± 0.1 µg cm−2) and triterpenoids (2.1 ± 0.4 µg cm−2), accompanied by minor quantities of alkanediols (0.2 ± 0.01 µg cm−2) and unidentified compounds (0.6 ± 0.3 µg cm−2).

Fig. 3 Compound class distribution in D. winteri leaf waxes. Coverages (μg/cm2) of each compound class in the waxes extracted with chloroform from the adaxial (top panel) and abaxial sides (bottom panel) of the D. winteri leaf were quantified by GC–FID. Averages ± SD (n = 5).

Chain length distribution of primary alcohols and esters in D. winteri leaf waxes

For a detailed analysis of chain length and isomer distributions in several compound classes, the D. winteri leaf waxes were purified using TLC (Supplementary Fig. S5). The least polar TLC fraction (Rf 1.0) of the wax mixtures extracted from both sides of D. winteri leaves contained very small quantities of alkanes (not detected in respective total wax mixtures), with similar chain length distributions in the adaxial and abaxial waxes (Supplementary Fig. S6). The adaxial wax fraction also comprised alkyl esters with overall chain lengths (including acid and alcohol moieties) ranging from C28 to C48 and fairly even distribution peaking at C38 (Fig. 4A). Each wax ester homolog consisted of a series of isomers with varying chain lengths of its acid and primary alcohol constituents (Supplementary Fig. S7A, B). For example, the C42 ester isomers were formed by C14–C24 acids and C18–C28 primary alcohols. The C30 and C32 as well as the C38 and C40 esters consisted mainly of C24 primary alcohol, while the C34, C36 and C42 esters comprised relatively large percentages of C28 alcohol. The prominent esters (C36–C42) all contained mainly C14 and/or C16 acid, whereas the longer esters (C44–C50) comprised relatively large amounts of C22 and C24 acids. In contrast, the shorter esters (C28–C34) mainly contained the C6 and C8 acids rarely seen in waxes. Summing across all ester homologs, the esterified acids had a broad chain length profile (Supplementary Fig. S7C), with C14 and C16 acids dominating (35% and 34%, respectively) and approximately equal amounts of C8, C20, C22 and C24 acids (6.4–7.8%). Across all ester homologs, the esterified alcohols showed a bimodal chain length distribution (Supplementary Fig. S7D) peaking at C22/24 (25%/44%) and C28 (16%).

Fig. 4 Chain length distribution in different wax compound classes of D. winteri leaf waxes. Relative amounts of each compound percentage of compound class), with total carbon number of each homolog or functional group position of each isomer given along the x-axis. Panels with bars pointing up or down show adaxial or abaxial wax compositions, respectively, for (A) esters, (B) primary alcohols, (C) secondary alcohols and (D) diols. Chain length distributions in (A, B, C) as well as diol isomer distributions in (D) were determined by GC–FID. Amounts of isomers within each secondary alcohol homolog in (C) were quantified by GC–MS using characteristic α-fragments. Averages ± SD (n = 3).

The wax from the adaxial leaf surface yielded a fraction migrating together with a primary alcohol standard (Rf 0.32) and containing C22 to C30 wax alcohols. The homologous series had a characteristic bimodal distribution (Fig. 4B), with C28 and C24 alcohols dominating (40% and 27%, respectively) and the C22, C26 and C30 alcohols contributing roughly equal amounts (7.0%, 10% and 14%, respectively). The corresponding fraction of the abaxial wax comprised mainly C28 alcohol (70%), along with smaller amounts of the C22, C24, C26 and C30 homologs (0.6%, 6.7%, 17% and 5.0%, respectively). Thus, the chain length distributions of wax primary alcohols differed between both leaf sides most pronouncedly in the relative amounts of the C24 homolog.

Composition of secondary alcohols and alkanediols in the waxes covering the adaxial and abaxial sides of the D. winteri leaf

The most prominent fraction (Rf 0.62) in the waxes from both leaf sides co-migrated with a secondary alcohol standard and was therefore suspected to contain respective wax alcohols. The major compound in this fraction was identified as nonacosan-10-ol based on comparisons of its MS fragmentation pattern with the nonacosan-10-ol spectrum in the literature (Simó et al. 1991) and a standard (Supplementary Fig. S8). Further compounds in the same fractions were identified as homologous C27, C31 and C33 secondary alcohols, also based on their MS characteristics.

GC–FID analysis showed that the C29 secondary alcohol accounted for approximately 98% and 95% of the TLC fraction in the waxes extracted, respectively, from the adaxial and abaxial sides of the D. winteri leaf. The adaxial leaf wax fraction further comprised 0.27% C27 alcohol, 0.17% C28 alcohol, 0.39% C30 alcohol, 0.77% C30 alcohol and trace amounts of C33 alcohol (Fig. 4C). The corresponding fraction of the mixture from the abaxial side contained 0.45%, 0.15%, 0.48%, 3.4% and 0.05% of the same secondary alcohol homologs.

Because secondary alcohol homologs with even carbon numbers were of special interest for deducing biosynthetic pathways, the secondary alcohol fraction was investigated in more detail using GC–MS. The fraction of the adaxial wax mixture showed a GC peak between the C27 and C29 secondary alcohol peaks, and thus likely representing a C28 secondary alcohol (Fig. 5A). It had MS fragments characteristic of alcohols along with ions m/z 467 ([M]+) and m/z 482 ([M-15]+), together confirming the C28 alcohol structure (Fig. 5B). Within the GC peak, two C28 alcohols with hydroxyls on C-9 and C-10 could be distinguished based on characteristic pairs of α-fragments (of respective trimethylsilyl (TMS)-ether derivatives) m/z 215 ([C9H18OSi(CH3)3]+) and m/z 369 ([C20H40OSi(CH3)3]+) versus m/z 229 ([C10H20OSi(CH3)3]+) and m/z 355 ([C19H38OSi(CH3)3]+). The two isomers could not be base-line separated under our GC conditions, but selected-ion traces showed that the C-10 isomer eluted slightly before the C-11 isomer (while also confirming the pairwise assignment of α-fragments; Fig. 5C). The GC elution of both the peak comprising both isomers very close to the half-way point between the C27 and C29 secondary alcohol peaks shows that all three homologs very likely have unbranched structures. In the same fraction of the adaxial wax mixture, a peak eluting between the C29 and C31 secondary alcohols showed ions m/z 511 ([M]+), and m/z 495 ([M-15]+) along with two pairs of α-fragments m/z 229 ([C10H20OSi(CH3)3]+) and m/z 383 ([C21H42OSi(CH3)3]+) as well as m/z 243 ([C11H22OSi(CH3)3]+) and m/z 369 ([C20H40OSi(CH3)3]+) (Fig. 5D). All this evidence taken together identified two isomeric C30 secondary alcohols with hydroxyl groups on C-10 and C-11. The C-10 alcohol isomer eluted slightly before the C-11 isomer (Fig. 5E). The same C28 and C30 secondary alcohol isomers were also detected in the corresponding fraction of wax extracted from the abaxial side of the D. winteri leaf (data not shown).

Fig. 5 GC–MS identification of secondary alcohols with even carbon numbers in the adaxial surface wax of D. winteri leaves. (A) Selected-ion chromatogram (m/z 73) of the TLC fraction containing secondary alcohols. (B) Mass spectrum and fragmentation diagram of the TMS-ether of the C28 secondary alcohol peak comprising isomers with C-9 and C-10 hydroxyls. (C) Selected-ion chromatograms of the α-fragments characterizing C28 9-ol and C28 10-ol. (D) Mass spectrum and fragmentation diagram of the TMS-ether of the C30 secondary alcohol peak comprising isomers with C-10 and C-11 hydroxyls. (E) Selected-ion chromatograms of the α-fragments characterizing C30 10-ol and C30 11-ol.

Further inspection of the α-fragments in mass spectra of each secondary alcohol homolog peak in the TLC fraction enabled the quantification of the positional isomers involved. In the C29 secondary alcohol of the adaxial wax, only one isomer, nonacosan-10-ol, could be detected (Fig. 4C). In contrast, the C27 homolog comprised two isomers with hydroxyls on C-10 and C-8 in a ratio of 9:1. Similarly, the C28 alcohols were a 3:1 mixture of the C-10 and C-9 isomers, the C30 alcohols were a 6:1 mixture of C-10 and C-11 isomers, and the C31 alcohols were a 3:1 mixture of the C-10 and C-12 isomers. The corresponding fraction from wax extracted from the abaxial leaf side showed similar isomer ratios. Overall, the D. winteri leaf waxes had secondary alcohols with hydroxyl groups mainly on C-10 or other even-numbered carbon atoms.

In a relatively polar TLC fraction (Rf 0.24), a range of alkanediols were identified by GC–MS. Diols with chain length C29 strongly dominated, and all isomers had one hydroxyl group on C-10 and the other one in varying positions between C-3 and C-15 (Fig. 4D). The diol fraction of the adaxial wax mixture comprised mainly nonacosane-4,10-diol and nonacosane-7,10-diol (38% and 33%, respectively), together with nonacosane-5,10-diol, nonacosane-10,13-diol, nonacosane-3,10-diol and nonacosane-6,10-diol (19%. 6.9%, 2.1% and 0.9%, respectively) as well as the C27 homolog heptacosane-4,10-diol (0.5%). In the corresponding fraction of abaxial wax, nonacosane-4,10-diol (58%) was accompanied by nonacosane-5,10-diol (19%), nonacosane-10,14-diol (3.3%), nonacosane-10,15-diol (15%) and small amounts of nonacosane-3,10-diol, nonacosane-6,10-diol and nonacosane-7,10-diol (1.5%, 1.6% and 1.4%, respectively), together with nonacosane-10,13-diol (0.3%). Thus, the two sides of the D. winteri leaf differed markedly in their wax alkanediol compositions, with nonacosane-7,10- and -10,13-diols restricted mainly to the adaxial surface, and nonacosane-10,14- and -10,15-diols to the abaxial side. In contrast, nonacosane-3,10-, -4,10-, -5,10- and -6,10-diols were present on both sides of the leaf in similar proportions.

Identification of ketols in D. winteri leaf wax

A moderately polar TLC fraction (Rf 0.25) from the abaxial wax of D. winteri leaves was suspected to comprise compounds with secondary functional groups. Four compounds in this fraction showed MS fragments m/z 73, 103 and 129 (Fig. 6A), indicating the presence of a single hydroxyl group (Ubik et al. 1975, Holloway et al. 1976). They were accompanied by an ion m/z 119 characteristic of compounds with non-vicinal secondary hydroxyl and keto groups (Jetter and Riederer 2000). All four compounds exhibited further prominent fragments m/z 496 and small amounts of ion m/z 510 (Fig. 6B) interpreted as respective [M-15]+ and [M]+ of C29 ketol structures (in their mono-TMS ether derivative form). Compounds 1–3 had pairs of α-fragments m/z 229/383 and m/z 243/369 indicating a hydroxyl group on C-10 of the C29 hydrocarbon chain, albeit with additional carbonyls in either the short or the long alkyl tails. Further smaller α-fragments located these carbonyls on C-15, C-5 and C-4, and fragments m/z 157 ([C5H8OSi(CH3)3]+) and 171 ([C6H10OSi(CH3)3]+) differentiated the C-4 and C-5 ketols (Jetter and Riederer 1996). Based on all this evidence, compounds 1–3 were thus identified as 10-hydroxynonacosan-4-one, 10-hydroxynonacosan-5-one and 10-hydroxynonacosan-15-one, respectively. Compound 4 had a pair of α-fragments m/z 299 and 387 characterizing a hydroxyl group on C-15, and further smaller α-fragments that indicated a carbonyl group on C-10 (Fig. 6B), together identifying it as 15-hydroxynonacosan-10-one. Reduction of the four compounds with lithium aluminum hydride (LAH) yielded three C29 alkanediols with hydroxyls on C-4/10, C-5/10 and C-10/15, thus corroborating the identification of the ketol isomers (Supplementary Fig. S9A, B). A fifth ketol may have been present in the same fraction, as indicated by a GC peak eluting shortly after the other ketols and also characterized by the ketol-specific ion m/z 119; however, its exact structure could not be assigned.

Fig. 6 GC–MS identification of ketols in the abaxial surface wax of D. winteri leaves. (A) Selected-ion chromatograms of the TLC fraction containing ketols, showing the retention time segment in which C29 ketols are eluting. (B) Mass spectra and fragmentation diagrams of the TMS-ethers of 10-hydroxynonacosan-4-one (compound 1), 10-hydroxynonacosan-5-one (compound 2) and a mixture of 10-hydroxynonacosan-15-one (compound 3) and 15-hydroxynonacosan-10-one (compound 4).

The relative amounts of the three ketols were assessed by integration of GC–MS traces of their characteristic α-fragments. The C29 4,10-ketol constituted 51.3% of the ketol fraction, along with 36.3% of the 5,10-ketol and 12.4% of the 10,15-ketol (Supplementary Fig. S9A). These compounds were present in trace quantities, and thus the absolute value could not be determined. The corresponding C-4/10, C-5/10 and C-10/15 diols resulting from LAH reduction had a very similar isomer distribution to the ketol isomers detected (Supplementary Fig. S9B). In the corresponding TLC fraction of the wax extracted from the adaxial surface of the D. winteri leaf, no ketols could be detected.

Identification of novel alkanetriols in D. winteri leaf wax

A relatively polar TLC fraction (Rf 0.1; migrating together with a fatty acid standard) of the wax extracted from the abaxial side of D. winteri leaves contained compounds that had not been described in plant waxes before. GC–MS analysis of TMS derivatives revealed five compounds, designated as 5–9, that were base-line separated under the present GC conditions (Fig. 7A). All five compounds showed MS fragments m/z 73, 103 and 129 expected for all wax alcohol TMS ethers as well as the ion m/z 147 indicating the presence of multiple hydroxyl groups (Eglinton et al. 1968, Busta et al. 2018). The compounds also showed combinations of low-abundance MS ions m/z 672 ([C29H57(OSi(CH3)3)3]+), m/z 657 ([M-CH3]+) and m/z 582 ([M-HOSi(CH3)3]+) suggesting isomeric C29 alkanetriol structures (Fig. 7B).

Fig. 7 GC–MS identification of alkanetriols in the abaxial surface wax of D. winteri leaves. (A) Selected-ion chromatogram (m/z 73) of the TLC fraction containing triols, showing the retention time segment in which C29 triols are eluting. (B) Mass spectra and fragmentation diagrams of the tris-TMS ethers of nonacosane-7,10,14-triol (compound 5), nonacosane-7,10,15-triol (compound 6), nonacosane-4,7,10-triol (compound 7), nonacosane-5,10,15-triol (compound 8) and nonacosane-4,10,15-triol (compound 9). The peak on the left of peak 1 is C30 primary alcohol.

Compound 9 had three pairs of α-fragments (Supplementary Figs. 5, 7B), respectively, comprising one TMS group (m/z 299 [C15H30OSi(CH3)3]+; m/z 145 [C4H8OSi(CH3)3]+), two TMS groups (m/z 317 [C10H19(OSi(CH3)3)2]+; m/z 457 [C20H39(OSi(CH3)3)2]+) and three TMS groups (m/z 475 [C15H28(OSi(CH3)3)3]+; m/z 629 [C26H50(OSi(CH3)3)3]+). The α-fragments containing two TMS groups were accompanied by daughter ions generated by loss of TMSOH (Δm/z −90), and those containing three TMS groups showed loss of one or two TMSOH units (Δm/z −90 and Δm/z −180). Based on respective fragment masses, the position of all three hydroxyl groups could be assigned unambiguously, and compound 9 was thus identified as nonacosane-4,10,15-triol.

Compounds 5–8 had MS fragmentation patterns also including three prominent pairs of α-fragments and respective elimination products Δm/z −90 and Δm/z −180, suggesting that they were positional isomers of 9. The specific fragment masses for each compound allowed assignment of the hydroxyl positions, such that compounds 5–8 were identified as nonacosane-7,10,14-triol, nonacosane-7,10,15-triol, nonacosane-4,7,10-triol and nonacosane-5,10,15-triol (Fig. 7B). Integration of the GC–MS data revealed a relatively even distribution of the newly identified triols, with nonacosane-4,7,10-triol as the major isomer and nonacosane-7,10,14-triol present in relatively small amounts (Fig. 7A). The triols could not be detected in the total wax mixture, so their absolute amounts could not be determined, and they must be regarded as trace components. No triols could be detected in the corresponding TLC fraction of the wax extracted from the adaxial surface of the D. winteri leaf.

Discussion

Our investigations of the D. winteri leaf surfaces showed that (i) the entire abaxial side of the leaf (including stomatal pores) was lined with a reticulate material bearing wax crystals, (ii) shapes and arrangements of epicuticular wax crystals differed between both leaf sides, (iii) wax mixtures on both sides of the leaf were dominated by secondary alcohols with C-10 hydroxyls, (iv) diol and triol isomers were unevenly distributed on both sides of the leaf, and (v) overall wax compositions differed between both leaf surfaces.

Investigation of the reticulate materials lining the abaxial surface of the D. winteri leaf

Both leaf surfaces of D. winteri were covered by a network of tubule-shaped wax crystals that were readily removed by surface washes with chloroform, an organic solvent known to dissolve wax almost instantaneously and quantitatively (Riederer and Schneider 1989). After repeat solvent washes, the adaxial surface appeared microscopically smooth, similar to many other plant epidermis surfaces investigated by SEM after solvent treatment, and likely showing the outside of the cutin matrix (Ganeva et al. 2015). In contrast, the abaxial surface of D. winteri leaves after wax removal showed an unusual reticulate structure that had been described before but not characterized. Here, we found that reagents known to degrade lipid polyesters decompose this structure, thus confirming earlier speculations that it consists of cutin (Bongers 1973, Feild et al. 1998). It seems plausible that the three-dimensional arrangement of the cutin matrix seen after wax removal existed in similar form on the native leaf surface, implying that the structure of the polymer comprised substantial micron-scale cavities. It will be interesting to investigate what special factors dictate the formation of this characteristic cutin structure.

Stomata were detected only on the abaxial side of the leaf, both by light microscopic inspection of the intact tissue and by SEM after treating the abaxial surface with adhesive tape. In contrast, SEM investigation of the native surface revealed only shallow humps of wax crystals in lieu of stomata, consisting of nanotubules similar to those on pavement cells but in denser arrangements. Based on the lateral dimensions of the humps, it is likely that they span the entire surface area of guard cells as well as the stomatal pores. The cutin meshwork associated with the waxes on the stomatal humps appeared slightly more porous than that lining the pavement cells. All this evidence distinguishes the stomatal plugs of D. winteri from those of gymnosperms, which consist only of wax and not of cutin (Jeffree 2006, Wen et al. 2006). The wax plugs of most gymnosperm species also do not fill the entire stomatal antechamber, and their stomata thus mostly appear as depressions of the overall surface rather than humps protruding from it.

Taken together, the chemical and morphological differences between the stomatal plugs of D. winteri and gymnosperms suggest different biological functions in both plant systems. Our results support the earlier hypothesis that stomatal plugs serve to keep the pores dry (Brodribb and Hill 1997, Mohammadian et al. 2009), where convex, hydrophobic humps may help shed water droplets. It is interesting to further speculate that the cutin reinforcement of the D. winteri plugs stabilizes them against erosion, which is likely to affect the exposed humps more than the sunken wax plugs of gymnosperms. Overall, our findings thus underscore the importance of stomatal humps for species like D. winteri growing in humid cloud forest habitats (Smith 1945). This is in contrast to many conifers adapted to arid climates, where stomatal plugs contribute to drought resistance (Jeffree et al. 1971, Brodribb and Hill 1997, Šantrůček 2022).

The material left after cutin removal had a structure also clearly differing between the stomatal plugs and the adjacent pavement cells. Based on the chemical treatment we employed and the general understanding of epidermal surface structures, this material most likely consists of carbohydrates, as part of the subtending cell walls. Overall, our experiments thus revealed structural differences between cell types in all three cell wall elements, including carbohydrates, cutin and waxes, pointing to differences in all the biosynthesis machineries involved and specific regulation of them in guard cells.

Wax crystal variation across the D. winteri leaf

The wax crystals on the adaxial side of the D. winteri leaf surface had varying shapes, including nanotubules and planar or curved structures with wavy edges. It is plausible that the nanotubules involved are formed, at least in part, by the nonacosan-10-ol and diols present in the wax mixture. Accordingly, the presence of much fewer nanotubules on the adaxial surface may be explained by the substantially lower absolute and relative amounts of both secondary alcohol and diols in the abaxial wax mixture compared with its adaxial counterpart. Conversely, the higher percentages of primary alcohols, esters and triterpenoids in the adaxial wax may lead to the formation of further crystals with shapes deviating from the characteristic nanotubules. In particular, primary alcohols have previously been associated with planar crystal forms (Jetter et al. 2006), and the crystals on the adaxial surface of the D. winteri leaf may therefore be formed by mixtures of primary and secondary alcohols. Interestingly, similar variation of tubular and variously curved wax crystal shapes had been reported for Papaver organ surfaces with wax mixtures comprising secondary and primary alcohol percentages similar to those on the adaxial side of the D. winteri leaf.

Our chemical analyses showed that the wax mixture extracted from the abaxial side of the D. winteri leaf accumulated substantially more wax, strongly dominated by nonacosan-10-ol and its diol derivatives. This finding confirms numerous previous reports on diverse species, all associating epicuticular wax nanotubules with similar dimensions and proportions with this specific secondary alcohol (Barthlott et al. 1998). Initially, in vitro reconstitution of native wax mixtures containing high amounts of nonacosan-10-ol (Jeffree et al. 1975) or purified nonacosan-10-ol (Jetter and Riederer 1994) showed that it is able to form tubular aggregates. Later, direct analyses of native wax nanotubules confirmed that they consist primarily of this single wax component (Wen et al. 2006). Based on all the evidence, it is very likely that the wax nanotubules on the abaxial surface of the D. winteri leaf are also formed by nonacosan-10-ol and corresponding nonacosanediols.

The wax nanotubules on pavement cells of the abaxial side of D. winteri leaves appeared distinctly longer than those on neighboring guard cells, pointing to possible differences in crystal compositions between both cell types. Comparable differences in wax nanotubule dimensions had been reported for other species, in comparisons of different tissues. Interestingly, the adaxial surface of Taxus baccata needles had relatively long tubules and a 3:1 ratio of secondary alcohol /diols, and shorter tubules and a 1:4 ratio on the abaxial surface (Wen et al. 2006). Similar differences in wax tubule lengths were reported for both sides of the Nelumbo nucifera leaf, associated with similar differences in alcohol/diol ratios (Ensikat et al. 2011). It may therefore be speculated that the relative amounts of secondary alcohol and diols in a wax mixture will affect nanotubule dimensions, and different alcohol/diol ratios may cause different appearances of the wax crystals on pavement and guard cells lining the abaxial side of the D. winteri leaf.

The different lengths and likely different compositions of wax crystals on pavement and guard cells must be due to differences in the wax biosynthesis machineries of both cell types. Our findings thus imply autonomous wax biosynthesis in guard cells, and they make lateral diffusion of waxes between neighboring cells improbable. This conclusion confirms previous circumstantial evidence for other species, where, for example, guard cells on Arabidopsis stems likely lacked characteristic secondary alcohols and ketones (Greer et al. 2007), while sunken stomata on gymnosperm needles had distinctly more wax than neighboring pavement cells (Wen et al. 2006).

Secondary alcohol composition and biosynthesis

The chain length and isomer distributions of secondary alcohols in D. winteri leaf wax shed light on the mechanisms underlying their formation. The predominant C29 alcohol had the hydroxyl group exclusively on C-10, while the C27, C31 and C33 alcohols comprised mixtures of isomers with C-8, C-10 or C-12 hydroxyls but no C-9 or C-11 hydroxyls. The secondary alcohols, thus, had a characteristic pattern of odd-numbered homologs with hydroxyls restricted to alternating, even-numbered carbons. Similar distributions had previously been reported for wax secondary alcohols of Papaveraceae, Ranunculaceae, Berberidaceae and Nelumbonaceae species (Holloway et al. 1976, Barthlott and Theisen 1994, Jetter and Riederer 1996, Koch et al. 2006, Tomaszewski and Zieliński 2014), implicating common and fairly wide-spread biosynthetic mechanisms. Accordingly, it had been concluded before that the characteristic positions of functional groups on every other carbon result from Claisen condensation reactions that may be catalyzed either by KAS/KCS enzymes involved in fatty acid elongation or by PKS enzymes (Busta and Jetter 2018). It had also been suggested that the 3-keto groups of the condensation reactions may, in later pathway steps, be converted into the hydroxyls of wax secondary alcohols (Busta and Jetter 2018). Our analyses of D. winteri leaf waxes now confirm the characteristic distribution of secondary alcohol isomers, and thus biosynthesis involving condensation reactions that establish the functional groups before the hydrocarbon chain is fully formed. This is in contrast to previous results for A. thaliana, where hydroxyls are introduced into pre-formed hydrocarbon structures. There, the P450-dependent monooxygenase MAH1 hydroxylates alkane substrates to form secondary alcohols with hydroxyls on even- and odd-numbered carbons (Wen and Jetter 2009).

The homolog and isomer patterns found in D. winteri leaf wax allow further insights into the mechanisms installing the secondary hydroxyls during hydrocarbon chain growth. Thus, the major wax component, nonacosan-10-ol, may be formed via C12 or C22 elongation intermediates with 3-ketoacyl or 3-hydroxyacyl structure (Fig. 8). They may be elongated to corresponding C30 hydroxy/ketoacyl-CoAs that can be decarboxylated (or reduced and decarbonylated) into nonacosan-10-ol. Alternatively, they may be linked with fatty acyl-CoAs by decarboxylative head-to-head condensation reactions into C29 ketols or diketones, which are then transformed into the secondary alcohol. By analogy, the C27, C31 and C33 secondary alcohol isomers may be biosynthesized from C10/12/14 or C20/22/24 3-hydroxy/ketoacyl precursors. Finally, the secondary alcohols with even-numbered carbon chain must be formed by the same mechanism as their odd-numbered homologs, but involving precursors with odd carbon numbers. Accordingly, head-to-head condensation of odd-numbered 3-hydroxy/ketoacyls or odd-numbered acyl-CoAs can explain all the even-numbered secondary alcohols detected in D. winteri waxes. Alternatively, these alcohols may also be formed by elongation of odd-numbered 3-hydroxy/ketoacyls but would require unlikely combinations of C21/23 and C11/13 precursors that would need to be derived from two separate (plastidial and ER-bound) pathways. Overall, our detailed homolog and isomer analyses thus lead us to hypothesize head-to-head condensation mechanisms for the formation of wax secondary alcohols, and it will be interesting to test this experimentally.

Fig. 8 Proposed biosynthetic pathways forming nonacosan-10-ol, diols and triols for leaf wax of D. winteri. Different colors of reaction arrows indicate P450 enzymes with different regio-specificities (arctic blue, hydroxylation on C-4 and C-5; cobalt blue, hydroxylation on C-7 and C-13; ruddy blue, hydroxylation on C-14 and C-15). The red ‘X’ indicates that P450-catalyzed hydroxylation of C29 alkane to nonacosan-10-ol is unlikely.

Alkanediol, ketol and triol composition and biosynthesis

All the alkanediols identified in D. winteri leaf waxes had one hydroxyl located on C-10 and a second group in various positions on either side of C-10. This finding suggested that the diols were derivatives of nonacosan-10-ol, likely formed by addition of a hydroxyl group in specific positions along the hydrocarbon chain. Eight C29 diols found in the wax mixtures from both leaf surfaces had been reported before (Wen et al. 2006), and two novel structures were identified as C29 10,14-diol and C29 10,15-diol. Interestingly, the latter two compounds are homologs of the C31 12,16-diol and C31 12,17-diol reported from Myricaria germanica leaf wax, with identical constellations of two hydroxyls, respectively, four and five carbons apart. Overall, the diols found in D. winteri leaf waxes resemble those accompanying nonacosan-10-ol in the surface wax mixtures of diverse other species, most notably in the Ranunculales (Jetter and Riederer 1996), Proteales (Barthlott et al. 1996), gymnosperms (Franich et al. 1978, Tulloch and Bergter 1981) and mosses (Neinhuis and Jetter 1995). Based on the structural similarity between nonacosan-10-ol and the diols, it had been suggested early on that the diols are biosynthetic derivatives of nonacosan-10-ol (Hunt and Baker 1979, Tulloch and Bergter 1981). It seems plausible that the D. winteri diols are also formed by hydroxylation of nonacosan-10-ol, with the initial hydroxyl possibly directing the relative position of the second hydroxyl group.

The mechanisms installing the second hydroxyl group can be gauged from their characteristic distributions across the two D. winteri leaf surfaces. First, the C29 10,14-diol and C29 10,15-diol were restricted to the abaxial side of the leaf, suggesting a hydroxylase enzyme expressed only in the epidermis on that side of the leaf. It is interesting to note that the two diols have hydroxyls in the same positions as the two major secondary alcohols found in Arabidopsis wax, C29 15-ol and C29 14-ol. The latter compounds are formed by hydroxylation of C29 alkane by the P450 enzyme MAH1, and an analogous hydroxylation of C29 secondary alcohol by an MAH1-like enzyme may yield the two D. winteri diols. Second, the C29 3,10-, 4,10-, 5,10- and 6,10-diols were found in similar proportions on both sides of the D. winteri leaf, implying that they may all be formed by the same enzyme. It seems plausible that this hydroxylase is also a P450 enzyme, with slightly different and overall lesser product specificity than MAH1. Third, the C29 7,10- and 10,13-diols accumulated mainly in the wax lining the adaxial leaf surface, suggesting that they are formed independently of the other diols. These two diols have characteristic hydroxyl constellations, with the second hydroxyl installed in the same distance on either side of C-10, and the hydroxylase involved may therefore have relatively strong positional specificity dictated by the initial hydroxyl rather than the distance to the alkyl terminus. There are no literature reports on enzymes with these characteristics, but it may be speculated that the hydroxylase involved is also a P450 enzyme. Similar reactions are known to be catalyzed by P450 enzymes, including the Arabidopsis CYP96B5 transforming wax alkanes into corresponding primary alcohols (Zhang et al. 2020) and CYP77A6 introducing mid-chain hydroxyls into fatty acid precursors of cutin (Li-Beisson et al. 2009). Overall, our analyses of the wax diols on both sides of the D. winteri leaf suggest the presence of at least three P450 enzymes installing hydroxyl groups in characteristic regions of C29 secondary alcohol precursors (Fig. 8).

Further to the alkanediols, we identified ketols and alkanetriols all characterized by one hydroxyl on C-10. The functional group positions and isomer distributions of these ketols and triols were identical to those of the accompanying diols, suggesting that all these compound classes are biosynthetically related. It seems likely that the hydroxylases installing hydroxyls in the alkanediols may catalyze further oxidation reactions, either on the same carbons to form ketols or on different carbons to yield the triols (Fig. 8). Overall, our results suggest at least three P450 enzymes acting alone and in tandem to hydroxylate both secondary alcohols and their diol derivatives.

Materials and Methods

Plant materials

Drimys winteri was continuously cultivated outdoors near the University of British Columbia, Vancouver, Canada. Mature leaves of approximately 8 cm length were harvested and for wax extraction and microscopy. Several leaves were pooled for each sample, and five independent samples taken for wax analysis. Three independent samples were taken for each microscopy experiment.

Wax extraction

To selectively extract total waxes from each side of the leaves, the intact mature leaves were placed on a flexible rubber mat. A glass cylinder with a 10-mm diameter was gently pressed onto the leaf surface, the cylinder was filled with approximately 1.5 ml of CHCl3 (Aldrich, 99%, 1% EtOH stabilizer) at room temperature, then the CHCl3 was agitated gently with a Pasteur glass pipette for 30 s. The CHCl3 was removed from the cylinder and transferred into a new glass tube, and the extraction steps were repeated three times and all extracts from the same leaf side were combined. Samples were discarded if the leaf surface was damaged or if CHCl3 leaked out under the glass cylinder. A known quantity of n-tetracosane was immediately added as an internal standard. The solvent was evaporated at 50°C under a stream of N2 (Praxair, >99.998%) and transferred to 2 ml GC autosampler vials with 250 μl inserts.

Thin layer chromatography

Total wax mixtures obtained by CHCl3 extraction from adaxial and abaxial sides of the leaves were individually loaded on silica plates (Merck) of 0.5 mm thickness with a concentrating zone. A mixture containing alkane, secondary alcohol, primary alcohol and fatty acid standards was loaded beside the wax sample on the plate. Plates were developed with mobile phase CHCl3:EtOH (99.5:0.5) using a sandwich technique (Tantisewie, et al., 1969), then air-dried and sprayed with primuline (Aldrich) dissolved in acetone and water (4:1, v/v). The bands were visualized under 365 nm UV light, scratched out, extracted with CHCl3 and analyzed with GC–MS.

Chemical analysis

Prior to GC analysis, all dried wax samples or TLC fractions were derivatized with 20 μl N,O-bis(trimethylsilyl)trifluoroacetamide (BSTFA, Aldrich, GC grade) and 20 μl pyridine (Aldrich, >99.8%, anhydrous) for 45 min at 70°C. Then, excess reagents were evaporated under N2, and samples were re-dissolved in CHCl3 (volume varying according to wax amounts).

Wax samples were analyzed as described previously (Buschhaus and Jetter 2012), using GC–MS to identify wax compounds and GC–FID for quantification. For compound identification, wax mixtures were separated by GC (6890N, Agilent Technologies) using an HP-1 capillary column (Agilent, 30 m length, 0.32 mm inner diameter, 0.1 µm film thickness) and cool-on-column injector, with oven temperature programmed for 2 min at 50°C, followed by 40°C min−1 to 200°C, held at 200°C for 2 min, increased 3°C min−1 to 320°C, held 30 min at 320°C. The inlet pressure was programmed for a constant 1.4 ml min−1 flow of helium carrier gas (Linde >99.995%). Compounds were identified by mass spectrometric detector (EI 70 eV, m/z 50–800, 1 scan s− 1, 5973N, Agilent Technologies). For compound quantification, wax mixtures were separated by GC under the same conditions as earlier, but with FID (HP ChemStation software package) and a constant flow of 2.0 ml min−1 of H2 carrier gas (Linde >99.995%). The absolute amount (µg) of each wax compound was established using the method established by Riederer and Schneider (1989) for comparison of GC peak areas with those of the internal standard. The extracted plant surface area was calculated based on the diameter of the cylinder opening. For the isomers that could not be separated by GC–FID, relative amounts (%) were determined by GC–MS averaging the quantities of the two characteristic hydroxyl α-fragments.

Scanning electron microscopy

Leaf samples were mounted on stubs using double-sided adhesive tape and air-dried for 1 week at room temperature. Then, specimens were sputter-coated with 5 nm of Ir using a Leica EM MED020 and visualized with a Helios NanoLab 650 Focused Ion Beam Scanning Electron Microscope at a 1.0-kV accelerating voltage and a working distance of 4.3 mm.

To localize stomata under the layer of epicuticular wax crystals, adhesive tape was attached to the abaxial surface of the leaf for 2 s and removed quickly. Then, the treated areas were cut out and mounted on stubs using double-sided adhesive tape and air-dried 1 week to further investigate by SEM as described earlier. To test whether microrelief structures on either side of the leaf consisted of wax, entire leaves were rinsed three times with warm CHCl3 for 30 s. Then, the leaf samples were dried and mounted with either side up on stubs using double-sided adhesive tape for SEM as described earlier. To further test whether micro-structures consisted of cutin, de-waxed samples were incubated in 0.5 m H2SO4 in methanol at 85°C for 4 h. Then, samples were dried and mounted on stubs using double-sided adhesive tape to further investigate by SEM as described earlier.

Light microscopy

Light micrographs were taken with an Olympus SZX10 stereomicroscope equipped with DP72 digital camera (Olympus) and an Olympus BX53 Light/Fluorescence Microscope equipped with DP80 dual chip color and monochromatic camera. Adaxial and abaxial sides of fresh leaves were visualized immediately after they were collected.

Supplementary Material

pcae053_Supp

Acknowledgments

The authors gratefully acknowledge technical assistance by Alberto Ruiz Orduna in early stages of the project, the University of British Columbia (UBC) BioImaging Facility for providing optical microscopes and assistance, and the UBC center for High-Throughput Phenogenomics for providing SEM and training.

Supplementary Data

Supplementary data are available at PCP online.

Data Availability

The data underlying this article will be shared upon reasonable request to the corresponding author.

Funding

Natural Sciences and Engineering Research Council (Canada) Discovery Grants Program (262461).

Author Contributions

Z.Z. and R.J. designed the project, Z.Z. acquired all data, D.M. analyzed ketols and Z.Z./R.J. all other data, Z.Z. wrote the draft and all authors edited the text.

Disclosures

The authors have no conflicts of interest to declare.
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