Present address: Science Research Center, Oceanography Section, Kochi University, Akebono-cho 2-5-1, Kochi-shi, Kochi, 780-8520, Japan.
Growth-dependent chemical and mechanical properties of cuticular membranes from leaves of Sonneratia alba
Article first published online: 13 FEB 2012
© 2012 Blackwell Publishing Ltd
Plant, Cell & Environment
Volume 35, Issue 7, pages 1201–1210, July 2012
How to Cite
TAKAHASHI, Y., TSUBAKI, S., SAKAMOTO, M., WATANABE, S. and AZUMA, J.-I. (2012), Growth-dependent chemical and mechanical properties of cuticular membranes from leaves of Sonneratia alba. Plant, Cell & Environment, 35: 1201–1210. doi: 10.1111/j.1365-3040.2012.02482.x
- Issue published online: 11 JUN 2012
- Article first published online: 13 FEB 2012
- Accepted manuscript online: 12 JAN 2012 09:10AM EST
- Received 16 July 2011; accepted for publication 1 January 2012
- cuticular membrane;
- growth dependence;
- Top of page
- MATERIALS AND METHODS
- Supporting Information
Chemical and mechanical properties of the leaf cuticular membranes (CMs) of a mangrove, Sonneratia alba J. Smith, were analysed at various leaf development stages to evaluate their tolerance to environmental stress. Our analyses demonstrate that the CMs from leaves of S. alba at different growth stages are generally rich in wax (21.5–25.7%) and cutin (52.4–63.4%) which rapidly accumulate at the early stages of leaf growth, while cutan (4.3–10.3%) and polysaccharide (2.3–7.7%) continuously accumulate throughout growth. Immature CMs are physically weak and highly viscoelastic. However, CMs become strengthened and stiffened during leaf expansion and maturation (by factors of about 1.5 and 2.4, respectively) while their flexibility decreases (68–83% decrease). Finally, the CMs lose their strength at the senescent stage (30–43% decreasement). Correlation analysis between chemical composition and mechanical properties revealed that the cutin matrix is mainly responsible for the high viscoelastic properties of CMs, while wax, cutan and polysaccharide contributed to their elasticity. Wax also affected the strength of the CMs, whereas cutan and polysaccharide showed rigidizing effect. Rapid accumulation of wax and cutin in the CMs after bud burst followed by the mechanical supports of cutan and polysaccharide in an isolateral manner contributed to the remarkable environmental tolerance of S. alba.
- Top of page
- MATERIALS AND METHODS
- Supporting Information
Sonneratia alba, which belongs to a genus of mangrove plants found in the Indo-West Pacific region, is an evergreen tree with tall conical pneumatophores arising from horizontal roots. Its leaves are opposite arranged, oval shaped, succulent and isolateral. They generally grow in the outermost area of the intertidal zone, and often form the seaward fringe (Tomlinson 1986). Their most striking feature is high tolerance against severe environmental growth conditions such as salinity, strong sunlight and salty wind (Wakushima, Kuraishi & Sakurai 1994). Highly developed surface layers are considered to play an important role in S. alba's durability.
Plant cuticular membranes (CMs) are continuous extracellular membranes located at the boundary of plant tissues and their external environment, especially on the aerial surface of leaves. CMs serve as the chemical and physical barrier against various biotic and abiotic impacts as well as mechanical support (Heredia 2003; Bargel et al. 2006; Stark & Tian 2006). In the case of S. alba, the primary role of the leaf CMs is providing physical protection against weathering, abrasions, leaching and water loss. CM is composed of a structural matrix called cutin, wax, polysaccharides and residual materials called cutan. Waxes are embedded inside the cutin and also cover the outer surface of CMs. Chemically, waxes are composed of complex mixtures of long chain hydrocarbons (alkanes, alcohols, ketones, fatty acids and esters), phenolics, terpenes and sterols (Tulloch 1976; Riederer & Markstädter 1996). Cutin, the major structural polymer of CMs, is an amorphous biopolyester formed by hydroxylated and/or epoxy-hydroxylated C16 and C18 fatty acids (Holloway 1980; Kolattukudy 1980; Jeffree 1996). Cutan is a structural polymer consisting of an aromatic fraction and/or an aliphatic part having polymethylenic chains (Domínguez, Heredia-Guerrero & Heredia 2011b) that is neither hydrolyzed with acid nor saponified with alkali (Nip et al. 1986; Jeffree 1996).
Mechanical properties of CMs have been extensively studied in tomato (Solanum lycopersicum L.) because of their importance in maintaining fruit quality (Bargel & Neinhuis 2005; Domínguez, Cuartero & Heredia 2011a). López-Casado et al. (2007) have reported the significant contribution of cuticular polysaccharides and cutin to elastic and viscoelastic properties, respectively, of tomato fruit CMs. Accumulation of flavonoids also contributes to stiffening of the cutin network in relation with fruit maturation (Domínguez et al. 2009). On the other hand, hydration of CMs increases their viscoelasticity and extensibility, suggesting a plasticizing effect of water on the cutin matrix (Petracek & Bukovac 1995; Wiedemann & Neinhuis 1998; Edelmann, Neinhuis & Bargel 2005). This kind of effect of water on cutin matrix was also obtained by atomic force microscopy (AFM) and nuclear magnetic resonance (NMR) spectroscopy at microscopic scale (Round et al. 2000).
Biomechanics of CMs of matured leaves of Yucca aloifolia L., Clusia fluminensis Planch. & Triana, Nerium oleander L. and Hedera helix L. were reported (Wiedemann & Neinhuis 1998; Edelmann et al. 2005). However, as pointed out by Domínguez et al. (2011a), little work has been done on the variations of the chemical and mechanical properties of CMs during growth. Maintaining the durability of the leaf CMs by increasing their physical strength is important for S. alba which grows in severe environmental conditions at the seaward fringe. Therefore, great efforts should be focused on understanding the process of biomechanical development of the S. alba leaf CMs during growth. In this paper, we describe the growth-dependent changes in chemical and mechanical properties of the CMs from leaves of S. alba, and discuss the chemical factors that contribute to their durability by analysing five pairs of leaves at different growth stages.
MATERIALS AND METHODS
- Top of page
- MATERIALS AND METHODS
- Supporting Information
Plant materials and isolation of CMs
Pairs of oppositely arranged isolateral leaves of S. alba J. Smith were collected from mature canopy trees which were sparsely distributed in the outer fringes of a mangrove forest in the Komi estuary (24°19′ N, 123°54′ E), Iriomote Island, Okinawa, Japan in October 2008, May 2009 and February 2010. Branches with five pairs of unfolded leaves were selected and all pairs of leaves were consecutively numbered from the top to the bottom (L1−L5) according to Lowman & Box (1983). The first pair of leaves (L1) from the branch top was at the immature expanding stage just after bud burst. The second leaf (L2) was also at the immature stage but almost attaining full expansion. The third (L3) and the fourth (L4) leaves were at the fully grown matured stage. The fifth leaf (L5) was at senescent stage before abscission. Thickness of the leaf was measured at 4–6 points per sheet using a digital micrometer (DP-1 VR, Mitutoyo Co., Tokyo, Japan). Surface areas of the leaves were calculated using image analysis software (ImageJ, National Institutes of Health, Bethesda, MD, USA).
Each pair of leaves was treated with an enzyme solution containing fungal pectinase (2%, w/v, Aspergillus niger, Sigma-Aldrich, St Louis, MO, USA) and a commercial cellulase preparation (0.5%, w/v, Meicelase CEP 16710, Trichoderma viride, carboxymethyl cellulase activity 2920 U, Meiji Seika Kaisha Ltd, Tokyo, Japan) in sodium acetate buffer (5 mm, pH 5.0) for 72 h at 36 °C. CMs from both adaxial (upper) and abaxial (lower) sides were separately isolated as thin films, washed with distilled water and dried. Surface of isolated CMs were morphologically characterized using low-voltage scanning electron microscopy (LV-SEM, VE-8800, Keyence Co., Osaka, Japan), at 500-fold magnification with 1.7 kV of accelerating voltage. Thickness of the CMs was measured in the same way as those of leaves.
Analyses of chemical constituents of CMs
The chemical components of the CMs were sequentially removed according to the modified method of Domínguez et al. (2009). Cuticular waxes were extracted three times with a mixture of chloroform/methanol (2:1, v/v) at 50 °C for 2 h. Dewaxed CMs were saponified three times with 1% potassium hydroxide in methanol at 70 °C for 2 h to estimate the cutin content. Decutinized residues were thoroughly washed with methanol after neutralization with 3% acetic acid in methanol, and then hydrolyzed according to the method of Saeman, Bubl & Harris (1945). The sample was mixed in 72% sulfuric acid, and kept at room temperature for 1 h, and subsequently the mixture was diluted with distilled water to 3% H2SO4, and autoclaved at 120 °C for 1 h. The final non-saponifiable and non-hydrolyzable residue was defined as cutan as described by Chen et al. (2008). Composition of monosaccharides obtained by acid hydrolysis of dewaxed and decutinized residues was determined by high performance anion-exchange chromatography (HPAEC, DX-500, Dionex, Sunnyvale, CA, USA) equipped with a pulsed amperometric detector and a column of CarboPac PA-1 (column size: 4.0 × 250 mm, Dionex) using 1.0 mm aqueous NaOH as an eluent at a flow rate of 1.0 mL min−1 as previously reported (Tsubaki et al. 2008). The polysaccharide content was quantified by the phenol-sulfuric acid method (Dubois et al. 1956) using a standard solution containing arabinose, xylose, rhamnose, galactose, glucose and mannose at a ratio determined by HPAEC. Removal of CM components by sequential extractions were evaluated by Fourier transform infrared (FT-IR) spectroscopy (FT/IR-4100, JASCO Co., Tokyo, Japan), at a 2.0 cm−1 resolution, and analytical range of 400–4000 cm−1 by the thin-film method or the KBr disc method.
Compositional analyses of cutin monomer
Depolymerization of cutin was carried out by hydrogenolysis or deuteriolysis according to the method of Walton & Kolattukudy (1972). Dewaxed CMs were refluxed in tetrahydrofuran with an excess amount (2.5 times by weight) of LiAlH4 or LiAlD4 (Tokyo Chemical Industry Co., Ltd, Tokyo, Japan) for 48 h at 70 °C. Reduced cutin monomers were extracted by diethyl ether after acidification by hydrochloric acid, then dehydrated by anhydrous sodium sulfate and evaporated to dryness under reduced pressure. Obtained samples were dissolved in excess N,O-bis(trimethylsilyl)-acetamide (Tokyo Chemical Industry Co., Ltd) and heated at 70 °C for 30 min to prepare trimethylsilyl (TMS) derivatives. Monomer composition of cutin was analysed by GC/MS (GC/MS 2010/PURVUM 2, Shimazu Co., Kyoto, Japan) equipped with a DB-1 column (J&W Scientific, 0.25 mm × 30 m, d.f. = 0.25 µm, Agilent Technologies, Inc., Santa Clara, CA, USA) with a helium carrier gas at a flow rate of 0.91 mL min−1. The column oven temperature was programmed from 195 to 240 °C at the rate of 2 °C min−1 and held for 10 min, then heated up to 300 °C at the rate of 10 °C min−1. The mass spectra were obtained in the range of 40–650 m/z by scanning mode with electron impact ionization at 70 eV.
Biomechanical analyses of CMs
Mechanical properties of the native and dewaxed CMs were measured by a tensile tester (Tack Tester TA-500, UBM Co., Kyoto, Japan), the equipment which is similar to that used by Wiedemann & Neinhuis (1998). Rectangular uniform segments 5 × 20 mm in size were cut out from the CM films, clipped with stainless clamps and deformed at a constant rate (0.01 mm s−1) at room temperature (23 °C). Data of strain and tensile force were collected twice per second. Elastic modulus (E, MPa) was obtained from a linear region at the initial part of the S-S curve. Breaking stress (σmax, MPa) and maximum strain (εmax, %) were determined at the point of failure of the CM films.
Multiple comparisons were performed by the Tukey–Kramer's HSD test at 5% level of significance, after one-way analysis of variance (anova). Confirmations of normality by Shapiro–Wilk test and rejections of outlier were carried out prior to data analyses. Correlation analyses were performed by using Pearson's product-moment correlation coefficient at 5 and 1% levels of significance.
- Top of page
- MATERIALS AND METHODS
- Supporting Information
Characterization of isolated CMs
Full leaves and enzymatically isolated CMs of S. alba were compared in respect to CMs physical and morphological characteristics at each stage of leaf growth. CMs of S. alba had similar surface morphologies on both sides of the leaves. As shown in Fig. 1, the CMs had a wrinkled surface, and stomata were observed in both adaxial and abaxial CMs. Table 1 shows summarized values of leaf area, weight per unit area and thickness of the leaves and isolated CMs. The values of average leaf area greatly increased with expansion of leaves from L1 (4.65 cm2) to L2 (17.35 cm2). The leaves at maturation and senescence stage (L3–L5) did not change in area size (P < 0.05). However, both values of the weight per unit area and thickness of the leaves still increased even after attaining full area size from 66.5 mg cm−2 (L1) to 133.3 mg cm−2 (L5) and from 530 µm (L1) to 1255 µm (L5), respectively.
|Area (cm2)†||4.65 ± 2.10a||17.35 ± 9.14b||17.77 ± 6.35b||20.86 ± 7.58b||19.31 ± 6.25b|
|Weight (mg cm−2)†||66.5 ± 7.2a||89.9 ± 10.1b||93.2 ± 20.4b||118.7 ± 22.4c||133.3 ± 29.2c|
|Thickness (µm)‡||530 ± 51a||720 ± 96b||840 ± 268c||1002 ± 340d||1255 ± 408e|
|Weight (µg cm−2)†||417 ± 109a||647 ± 169b||741 ± 37bc*||754 ± 127bc||888 ± 122c*|
|Thickness (µm)‡||3 ± 1a||9 ± 1b*||9 ± 2b*||9 ± 3b||10 ± 3b*|
|Weight (µg cm−2)†||457 ± 106a||682 ± 206b||692 ± 52b*||728 ± 86b||703 ± 131b*|
|Thickness (µm)‡||3 ± 1a||8 ± 2b*||7 ± 2b*||8 ± 2b||9 ± 2b*|
In the case of the CMs, the weight per unit area increased about 1.5-fold from L1 to L2; however, the increase reached a plateau during L3–L5. Rapid increase in the average thickness of the CMs was also observed from L1 to L2 (increment 5–6 µm), showing that both expansion and thickening of the S. alba leaf CMs take place mainly during growth from L1 to L2. No significant difference was observed between the weights of both sides of CMs (P < 0.05), while adaxial CMs tend to be slightly thicker than abaxial CMs.
FT-IR analysis of CM components
Validity of the solvent extractions for the sequential removal of the constituents of the CMs was spectroscopically verified by FT-IR prior to the compositional analyses. All CMs at different growth stages gave similar spectra. Figure 2 shows the representative data given by the adaxial CM at L4. The spectrum of the native CM had strong absorptions at 2923, 2851 and 1734 cm−1 assigned as asymmetric and symmetric stretching vibrations of methylene groups and stretching vibration of esterified carbonyl groups. These intense absorption bands showed predominance of cutin (Ramirez et al. 1992; Villena, Domínguez & Heredia 2000). Spectrum of the extracted waxes was characterized by a strong absorption at 1687 cm−1 and corresponded to the stretching vibrations of the carbonyl groups in addition to the strong alkyl peaks and a weak hydroxyl peak at 3349 cm−1. Removal of waxes from CMs slightly sharpened absorptions due to ester carbonyl, methylene and hydroxyl groups. Decutinization drastically weakened absorptions due to the methylene and ester carbonyl groups. Acid hydrolysis decreased the absorptions around 1000–1200 cm−1 together with lowering of hydroxyl group due to effective removal of polysaccharides (Kačurákováet al. 2000; Villena et al. 2000). Reappearance of the alkyl peaks in the final residue confirmed the presence of highly recalcitrant aliphatic polymer, cutan, in the S. alba leaf CMs. Broad absorption bands around 1650 cm−1 were assigned to C=C bonds (Villena et al. 1999; Chen et al. 2008) and presence of aromatic cores was indicated by an absorption at 1517 cm−1 due to C-C bonds conjugated with C=C bonds (Chen et al. 2008; Järvinen et al. 2011).
The chemical composition of the isolated leaf CMs of S. alba was gravimetrically analysed. The results are shown in Table 2 and Fig. 3. The overall quantitative results indicate presence of two sets of components that show different patterns of increase in concentration; one set of components was wax and cutin, and the other set was cutan and polysaccharide. The former were present at a low level in L1 and at a rather high levels throughout L2–L5 (Fig. 3a,b), while the latter gradually increased from L1 to L5 (Fig. 3c,d). The proportion of wax did not show any difference throughout L1–L5 (Table 2); however, its amount apparently increased from L1 (98–112 µg cm−2) to L2 (142–153 µg cm−2), with slight increase to L5 (156–168 µg cm−2): n.b. statistically significant difference was barely observed because of large variance (Fig. 3a). Cutin was the most abundant constituent followed by wax, and the sum of these two predominant components amounted to 74.2–88.5% of total CMs (Table 2). Similar to the wax content, the amount of cutin drastically increased from L1 (245–256 µg cm−2) to L2 (373–379 µg cm−2) although the values reached a plateau after L2 (Fig. 3b). These patterns of content increase of cutin and wax were consistent with the increase in the weight and thickness of the CMs (Table 1) showing that the construction of the main framework has completed during the rapid leaf expanding stages (L1 to L2). On the contrary, the amounts of cutan and polysaccharides continuously increased throughout the leaf expansion, maturation and aging stages (L1–L5). The content of cutan was in the range of 19–77 µg cm−2 (adaxial) and 23–66 µg cm−2 (abaxial) while the content of polysaccharides was in the range of 14–60 µg cm−2 (adaxial) and 13–49 µg cm−2 (abaxial). Accumulation of cutan and polysaccharide was evident at the matured and old senescent stages (Fig. 3c,d). Their contents were, however, significantly smaller than those of wax and cutin; 5.2–11.1% (cutan) and 3.3–7.5% (polysaccharides) (Table 2).
|Waxa||25.7 ± 8.2||23.1 ± 7.5||22.9 ± 3.9||24.1 ± 3.7||23.4 ± 4.4|
|Cutina||63.4 ± 10.1||60.1 ± 3.6||59.2 ± 6.4||57.5 ± 2.9||54.4 ± 4.3|
|Cutanb||5.0 ± 4.4||5.1 ± 2.3||6.5 ± 2.8||8.6 ± 4.2||10.2 ± 3.5|
|Polysaccharideb||3.5 ± 0.9||3.6 ± 0.6||5.8 ± 1.2||6.4 ± 1.3||7.4 ± 2.3|
|Arabinose||27.4 ± 2.4||31.9 ± 0.2||39.3 ± 1.7||37.1 ± 1.0||37.9 ± 2.5|
|Rhamnose||0.7 ± 0.4||0.6 ± 0.1||0.9 ± 0.6||0.1 ± 0||0.6 ± 0.2|
|Galactose||7.2 ± 0.5||4.7 ± 0.8||6.9 ± 0.8||7.5 ± 0.5||6.7 ± 0.7|
|Glucose||58.0 ± 4.5||54.1 ± 1.2||45.7 ± 1.9||45.8 ± 1.3||43.5 ± 2.5|
|Xylose||5.6 ± 1.7||5.8 ± 0.5||5.6 ± 1.6||6.8 ± 0.3||7.8 ± 0.8|
|Mannose||1.2 ± 0.3||2.8 ± 0.8||1.6 ± 0.6||2.6 ± 0.6||3.6 ± 0.1|
|Waxa||23.2 ± 4.0||21.5 ± 5.4||24.5 ± 5.4||24.5 ± 4.4||25.2 ± 4.7|
|Cutina||60.7 ± 11.8||60.9 ± 3.8||52.4 ± 5.5||55.9 ± 2.9||53.5 ± 3.3|
|Cutanb||4.3 ± 1.3||6.4 ± 2.4||8.4 ± 2.7||8.4 ± 2.7||10.3 ± 3.5|
|Polysaccharideb||2.3 ± 0.6||3.3 ± 0.4||4.4 ± 0.9||6.5 ± 1.3||7.7 ± 3.1|
|Arabinose||28.3 ± 4.4||27.3 ± 3.4||40.1 ± 0.9||39.4 ± 2.0||35.1 ± 0.4|
|Rhamnose||0.9 ± 0.3||0.8 ± 0.3||1.0 ± 0.5||0.3 ± 0.2||0.6 ± 0.2|
|Galactose||6.1 ± 1.1||8.8 ± 2.4||7.1 ± 1.2||8.0 ± 0.4||6.7 ± 0.4|
|Glucose||56.6 ± 3.5||47.8 ± 4.5||41.2 ± 0.9||41.9 ± 2.4||45.0 ± 1.1|
|Xylose||5.3 ± 1.2||11.1 ± 5.4||7.7 ± 0.1||7.6 ± 0.8||9.1 ± 0.7|
|Mannose||1.5 ± 0.5||4.2 ± 0.5||2.9 ± 0||2.8 ± 0.4||3.6 ± 0.2|
Polysaccharides contained in the CMs were hydrolyzed and their monosaccharide composition was determined by HPAEC as summarized in Table 2. Both adaxial and abaxial CMs showed similar monosaccharide compositions. Glucose was the most abundant monosaccharide amounting to 41.2–58.0%, followed by arabinose 27.3–40.1% with smaller contents of galactose (4.7–8.8%) and xylose (5.3–11.1%). The results presented indicate that the cuticular polysaccharides in the S. alba leaf CMs were composed of mixtures of cellulose, pectic and hemicellulosic polysaccharides.
Cutin monomer composition
Monomer composition of the cutin in the CMs was determined by reductive depolymerization followed by GC/MS analysis. As shown in Supporting Information Table S1 and Supporting Information Figs S1–S3, fragmentation patterns of the mass spectra (MS) were identified according to Walton & Kolattukudy (1972). All cutins from the S. alba leaf CMs were composed of at least 14 monomers of which 11 were identified as listed in Table 3. Predominant components were hexadecan-3-ol and octadecan-3-ol, accounting for about 20–25 and 30–55% of the total monomers, respectively. Combination analysis of the deuterium-labelled monomers allowed us to conclude that these components were derived from 9(10),16-dihydroxyhexadecanoic acid, which appeared as a single peak with a mixture of two isomers, and 9,10-epoxy-18-hydroxyoctadecanoic acid, respectively (Supporting Information Figs S2 & S3). Four monohydroxy alkans (hexadecan-1-ol, octadecan-1-ol, hexadecan-2-ol and heptadecan-3-ol) and three monohydroxy alkens (octadecen-2-ol, tetradecen-3-ol and octadecen-3-ol) were also detected as minor components, amounting to less than 5% in total. Both of adaxial and abaxial CMs had similar cutin monomer composition throughout the growth stages.
|Hexadecan-1-ol (C16:0)||0.23 ± 0.14||0.04 ± 0||0.18 ± 0.11||0.09 ± 0.01||0.25 ± 0.11|
|Octadecan-1-ol (C18:0)||0.12 ± 0.05||0.03 ± 0.02||0.03 ± 0.03||0.06 ± 0.02||0.13 ± 0.07|
|Hexadecan-2-ol (C16:0)||0.76 ± 0.09||0.57 ± 0.02||0.77 ± 0.17||0.72 ± 0.06||1.10 ± 0.23|
|Octadecen-2-ol (C18:1)||0.54 ± 0.18||0.71 ± 0.22||1.85 ± 0.20||1.22 ± 0.08||2.99 ± 0.10|
|Hexadecan-3-ol (C16:0)||23.70 ± 1.88||21.57 ± 0.84||25.57 ± 1.14||24.81 ± 0.27||20.91 ± 1.35|
|Tetradecen-3-ol (C14:1)||0.70 ± 0.08||0.48 ± 0.07||2.91 ± 0.18||1.70 ± 0.07||10.73 ± 0.56|
|Heptadecan-3-ol (C17:0)||0.18 ± 0.03||0.12 ± 0.01||0.27 ± 0.03||0.51 ± 0.02||0.14 ± 0.04|
|Unknown 1||2.96 ± 0.09||2.15 ± 0.13||0.20 ± 0.03||0.86 ± 0.06||0.42 ± 0.06|
|Octadecen-3-ol (C18:1)||0.56 ± 0.13||0.60 ± 0.05||0.79 ± 0.05||0.50 ± 0.04||3.33 ± 0.53|
|Octadecan-3-ol (C18:0)||54.02 ± 1.25||49.35 ± 0.45||47.48 ± 1.21||49.27 ± 0.28||31.71 ± 2.19|
|Unknown 2||–||0.79 ± 0.11||9.80 ± 0.08||4.71 ± 0.16||5.59 ± 1.62|
|Hexadecen-3-ol (C16:1)||0.90 ± 0.15||1.08 ± 0.09||3.29 ± 0.68||3.15 ± 0.10||11.75 ± 1.76|
|Octadecan-4-ol (C18:0)||11.57 ± 0.40||15.38 ± 0.60||2.75 ± 0.01||10.22 ± 0.07||3.38 ± 0.11|
|Unknown 3||3.76 ± 0.23||7.13 ± 0.69||4.11 ± 0.22||2.20 ± 0.07||7.56 ± 0.72|
|Hexadecan-1-ol (C16:0)||0.17 ± 0.01||0.08 ± 0.03||0.31 ± 0.09||0.10 ± 0.05||0.68 ± 0.08|
|Octadecan-1-ol (C18:0)||0.13 ± 0.02||0.03 ± 0.02||0.11 ± 0.04||0.04 ± 0.01||0.08 ± 0.01|
|Hexadecan-2-ol (C16:0)||0.62 ± 0.10||0.76 ± 0.06||0.84 ± 0.06||0.75 ± 0.11||1.08 ± 0.13|
|Octadecen-2-ol (C18:1)||0.86 ± 0.21||2.39 ± 0.17||1.73 ± 0.15||2.47 ± 0.15||4.61 ± 0.14|
|Hexadecan-3-ol (C16:0)||19.72 ± 0.63||20.23 ± 0.70||18.14 ± 0.64||20.84 ± 1.41||25.96 ± 0.88|
|Tetradecen-3-ol (C14:1)||3.08 ± 0.27||4.60 ± 0.50||3.31 ± 0.18||3.25 ± 0.33||4.28 ± 0.41|
|Heptadecan-3-ol (C17:0)||0.38 ± 0.15||0.18 ± 0.02||0.23 ± 0.01||0.19 ± 0.03||0.39 ± 0.02|
|Unknown 1||Tr||1.78 ± 0.07||0.24 ± 0.02||1.26 ± 0.04||0.94 ± 0.04|
|Octadecen-3-ol (C18:1)||0.70 ± 0.39||1.77 ± 0.12||1.60 ± 0.15||3.47 ± 0.09||4.06 ± 0.24|
|Octadecan-3-ol (C18:0)||57.36 ± 0.35||46.76 ± 0.96||47.87 ± 0.59||42.52 ± 0.35||37.02 ± 0.49|
|Unknown 2||–||7.47 ± 0.22||5.14 ± 0.11||7.32 ± 0.97||4.90 ± 0.28|
|Hexadecen-3-ol (C16:1)||5.70 ± 0.39||5.49 ± 0.19||7.84 ± 0.24||6.45 ± 0.80||6.21 ± 0.25|
|Octadecan-4-ol (C18:0)||3.90 ± 0.78||3.58 ± 0.14||4.12 ± 0.05||5.70 ± 0.19||5.94 ± 0.10|
|Unknown 3||6.96 ± 0.14||4.88 ± 0.26||8.52 ± 0.43||5.63 ± 0.37||3.85 ± 0.27|
Biomechanical properties of the isolated CMs were measured by the tensile test. Specimens from both adaxial and abaxial CMs gave similar profiles. Typical S-S curves, representative of both adaxial and abaxial CMs, are shown in Fig. 4. A clear biphasic (two phase with different slope) behaviour was observed in S-S curves from young leaf CMs (L1-L3), whereas most CMs from the fully matured leaves (L4 and L5) usually showed almost monophasic behaviour. Table 4 shows summarized values of biomechanical parameters of the isolated leaf CMs. The values in σmax increased with the leaf growth, attaining a maximum at L4 (7.29–8.74 MPa), then decreasing to 4.14–6.10 MPa at L5. The CMs isolated from immature leaves (L1) showed high extensibility with εmax of 7.55–9.07%; however, this value decreases during the leaf growth to 1.54–2.42% at L5. The E-values of the isolated CMs were below 200 MPa at L1 in both adaxial and abaxial CMs, but increased with leaf growth attaining values higher than 390 MPa in CMs from the fully matured leaves (L4 and L5). Additionally, both of the adaxial and abaxial CMs showed similar mechanical properties as expected from their similar morphological and chemical properties.
|σmax (MPa)||5.20 ± 1.31ac||6.51 ± 1.31ab||6.47 ± 1.99ab||7.29 ± 0.98b||4.14 ± 0.56c*|
|εmax (%)||9.07 ± 1.65a*||6.63 ± 1.22b||4.34 ± 1.96c||3.52 ± 1.06cd||1.54 ± 0.29d*|
|E (MPa)||169 ± 63a||279 ± 14b||303 ± 43b||398 ± 86c||403 ± 43c|
|σmax (MPa)||5.60 ± 1.52a||6.41 ± 0.80a||7.34 ± 1.37ab||8.74 ± 2.60b||6.10 ± 1.32a*|
|εmax (%)||7.55 ± 1.07a*||6.38 ± 0.95a||4.50 ± 1.55b||4.48 ± 1.17b||2.42 ± 0.46c*|
|E (MPa)||174 ± 66a||287 ± 71b||364 ± 48bc||441 ± 92c||410 ± 67c|
To investigate the role of wax in the mechanical properties of CMs, the tensile test was also applied to dewaxed CMs from the mature leaves (Fig. 4). In the adaxial side, the dewaxed CM at L4 showed 4.82 MPa of σmax, 5.43% of εmax and 162 MPa of E. The corresponding values of the abaxial side were 3.89 MPa, 4.16% and 138 MPa, respectively. By removal of waxes, both values of physical strength and stiffness of matured leaf CMs drastically decreased strongly, indicating that the accumulation of cuticular waxes in CMs contributes to the mechanical properties of CMs. After dewaxing and decutinization, all CMs became very brittle and did not maintain mechanical strength enough to carry out the tensile test.
Relationship between chemical and mechanical properties
The relationship between chemical and mechanical properties of the CMs was statistically evaluated and the results are summarized in Table 5. In the wax content, significant correlations were observed with the σmax value (R = 0.331) and the E-value (R = 0.433). The cutin content was positively correlated with the εmax value (R = 0.507) and negatively correlated with the E-value (R = −0.468). In contrast, the contents of cutan and polysaccharide showed good negative correlations with the εmax value (R = −0.549 and −0.694) and relatively weak positive correlation with the E-value (R = 0.237 and 392), respectively. No significant correlation was observed in the compositions of cutin monomers and monosaccharides.
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In the study presented here, we characterized the variation in chemical composition of the CMs from leaves of S. alba during expansion, maturation and aging stages. The quantitative analyses of the chemical constituents revealed a growth dependence (Fig. 3), consistent with previous developmental models of leaf CMs; deposition of wax and cutin rapidly proceeds in the early stages of development, whereas the quantity of cutan appears to increase at later stages (Jeffree 1996).
CMs of S. alba contained higher amount of wax (Table 2) than previously reported in H. helix, Olea europaea L., Agave americana L., Clivia miniata Reg. and Citrus limon Burm. f. These plants were shown to contain approximately 12, 26, 16, 7 and 8% of wax, respectively (Nip et al. 1986; Hauke & Schreiber1998; Johnson et al. 2007). By comparison, the leaf CMs of S. alba are rich in wax (Table 2). The abundance of the cuticular wax should provide higher water repellency to the leaves of S. alba which are often submerged in sea water and subjected to salty winds.
Overall similarity of chemical composition in the CMs from adaxial and abaxial sides confirmed the unique isolaterality of S. alba leaves (Tomlinson 1986). Both adaxial and abaxial CMs of S. alba leaves were found to have similar cutin monomer composition. The predominance of C18 monomers was in accord with monomer profiles of leaf CMs of H. helix, C. miniata, Hordeum vulgare L. and Camellia sinensis (L.) Kuntze (Riederer & Schönherr 1988; Richardson et al. 2007; Tsubaki et al. 2008; Graça & Lamosa 2010). Graça & Lamosa (2010) proposed that cutin has domains of linear structure when C18-epoxy monomers are dominant, whereas branched networks occur when the C16-dihydroxy monomers predominate. According to these criteria, linear structure might be predominated in the cutin of S. alba leaves.
Mechanical tests quantitatively showed how the leaves of S. alba acquired environmental tolerance. Physically weak and highly viscoelastic CMs at the immature stage were strengthened and stiffened during leaf growth together with the loss of high extensibility (Fig. 4; Table 4). However, physical strength of leaf CMs was finally lost at the senescent stage. Growth-dependent mechanical properties presented in this study can be attributed to the variation of the chemical constituents. As previously reported (Petracek & Bukovac 1995), removal of waxes resulted in decrease in the values of σmax and E, showing that wax has strengthening and stiffening effects on the CMs (Fig. 4).
The CMs isolated from different species or organs showed a wide diversity in chemical composition (Holloway 1980), and mechanical properties of CMs depend on their species of origin (Wiedemann & Neinhuis 1998). Yet most of the studies on biomechanics of CMs address fruit growth and ripening, and no similar work has been conducted on the alteration of mechanical properties of CMs during the growth of vegetative tissues (Domínguez et al. 2011a). The results on the growth-dependent chemical and mechanical properties presented here provide new information on the functional aspects of cuticular constituents in leaf epidermal layers of S. alba.
In the present study, the chemical composition of the CMs was statistically correlated with its mechanical properties (Table 5). Significant contribution of cuticular waxes to the CM biomechanics was in accord with previous suggestion made by Petracek & Bukovac (1995). Intra-cuticular waxes can be regarded as fillers that reduce free volume and restrict the segmental mobility of cutin molecules within the matrix (Petracek & Bukovac 1995; Bargel et al. 2006). Contributions of cutin to the increase in extensibility and decrease in stiffness and inverse contributions of polysaccharides demonstrated that the chemomechanic system, previously shown to operate in fruit CMs (López-Casado et al. 2007), also works in leaves. This stiffening and rigidizing effects of polysaccharides on the cutin matrix seemed to be reasonable as cellulose microfibrils probably stiffened cuticular matrix in the same way as in fibre-reinforced composite materials (Bargel et al. 2006; López-Casado et al. 2007). In addition, the accumulation of cutan during the leaf growth statistically correlated with decrease in extensibility and increase in stiffness of the CMs. Previously, cutan was suggested to be a factor that influenced the rigid appearance of the CMs (Bargel et al. 2006). Cutan present in the leaf CMs of S. alba contains C = C bonds and/or aromatic rings which weakens the stacking of molecular chains as evidenced by the FT-IR spectra (Fig. 1). We can conclude that cutan contributes to the rigidity of the CMs. These contributions of cuticular components may cause the modifications of the shape of the S-S curves (Fig. 4). The change of the S-S curves from biphasic to monophasic was in accord with leaf age. This might be due to uniform distribution of constituents, especially polysaccharides and cutan molecules in the CMs from matured leaves after formation of a rigid network structure; in turn, flexibility remains in the CMs from immature leaves where deposition of these constituents is still working.
The substantial weakening of CMs at L5 was not ascribed exclusively to the variations of the quantity of each constituent. As decomposition of constituents may occur during progress of senescence, the quality of the constituents may contribute to their declining.
In conclusion, the present study characterized growth-dependent chemical and mechanical properties of the CMs from S. alba leaves and further elucidated the chemical factors which influenced the biomechanics of the CMs. Although the amounts of wax and cutin achieved plateau at an early stage of leaf growth, deposition of cutan and polysaccharide continued throughout the growth. Physical strength and stiffness of the CMs increased with leaf expansion and maturation, but the strength weakened at the senescent stage. Flexibility of the CMs gradually decreased along with the leaf growth. The viscoelastic nature of the immature CMs was ascribed to cutin whereas the stiffening of the cutin matrix was owned by the other components, wax, cutan and polysaccharides. In addition, wax strengthens the cutin matrix. Cutan and polysaccharides showed a rigidizing effect. Isolateral development of CMs of S. alba leaves and rapid accumulation of wax and cutin in the CMs after bud burst followed by the mechanical supports of cutan and polysaccharides contribute to its remarkable environmental tolerance to allow growth at the seaward fringe.
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- 2005) Tomato (Lycopersicon esculentum Mill.) fruit growth and ripening as related to the biomechanical properties of fruit skin and isolated cuticle. Journal of Experimental Botany 56, 1049–1060. & (
- 2006) Structure-function relationships of the plant cuticle and cuticular waxes – a smart material? Functional Plant Biology 33, 893–910. , , & (
- 2008) Role of the extractable lipids and polymeric lipids in sorption of organic contaminants onto plant cuticles. Environmental Science and Technology 42, 1517–1523. , , , & (
- 2009) Biomechanics of isolated tomato (Solanum lycopersicum) fruit cuticles during ripening: the role of flavonoids. Functional Plant Biology 36, 613–620. , , , & (
- 2011a) An overview on plant cuticle biomechanics. Plant Science 181, 77–84. , & (
- 2011b) The biophysical design of plant cuticles: an overview. New Phytologist 189, 938–949. , & (
- 1956) Colorimetric method for determination of sugars and related substances. Analytical Chemistry 28, 350–356. , , , & (
- 2005) Influence of hydration and temperature on the rheological properties of plant cuticles and their impact on plant organ integrity. Journal of Plant Growth Regulation 24, 116–126. , & (
- 2010) Linear and branched poly(ω-hydroxyacid) esters in plant cutins. Journal of Agricultural and Food Chemistry 58, 9666–9674. & (
- 1998) Ontogenetic and seasonal development of wax composition and cuticular transpiration of ivy (Hedera helix L.) sun and shade leaves. Planta 207, 67–75. & (
- 2003) Biophysical and biochemical characteristics of cutin, a plant barrier biopolymer. Biochimica et Biophysica Acta – General Subjects 1620, 1–7. (
- 1980) The chemical constitution of plant cutins. In The Plant Cuticle (eds D.F. Cutler, K.L. Alvin & C.E. Price), pp. 45–85. Academic Press, London, UK. (
- 2011) Solid state 13C CP-MAS NMR and FT-IR spectroscopic analysis of cuticular fractions of berries and suberized membranes of potato. Journal of Food Composition and Analysis 24, 334–345. , , & (
- 1996) Structure and ontogeny of plant cuticles. In Plant Cuticles: An Integrated Functional Approach (ed. G. Kerstiens), pp. 33–82. BIOS Scientific Publishers Ltd, Oxford, UK. (
- 2007) Spectroscopic characterization of aliphatic moieties in four plant cuticles. Communications in Soil Science and Plant Analysis 38, 2461–2478. , , , & (
- 2000) FT-IR study of plant cell wall model compounds: pectic polysaccharides and hemicelluloses. Carbohydrate Polymers 43, 195–203. , , , & (
- 1980) Biopolyester membranes of plants: cutin and suberin. Science 208, 990–1000. (
- 2007) Biomechanics of isolated tomato (Solanum lycopersicum L.) fruit cuticles: the role of the cutin matrix and polysaccharides. Journal of Experimental Botany 58, 3875–3883. , , , & (
- 1983) Variation in leaf toughness and phenolic content among five species of Australian rain forest trees. Australian Journal of Ecology 8, 17–25. & (
- 1986) A new non-saponifiable highly aliphatic and resistant biopolymer in plant cuticles – evidence from pyrolysis and 13C-NMR analysis of present-day and fossil plants. Die Naturwissenschaften 73, 579–585. , , , & (
- 1995) Rheological properties of enzymatically isolated tomato fruit cuticle. Plant Physiology 109, 675–679. & (
- 1992) Fourier transform IR study of enzymatically isolated tomato fruit cuticular membrane. Biopolymers 32, 1425–1429. , , & (
- 2007) Cuticular permeance in relation to wax and cutin development along the growing barley (Hordeum vulgare) leaf. Planta 225, 1471–1481. , , , , , & (
- 1996) Cuticular waxes: a critical assessment of current knowledge. In Plant Cuticles: An Integrated Functional Approach (ed. G. Kerstiens), pp. 189–200. BIOS Scientific Publishers Ltd., Oxford, UK. & (
- 1988) Development of plant cuticles: fine structure and composition of Clivia miniata Reg. leaves. Planta 174, 127–138. & (
- 2000) The influence of water on the nanomechanical behavior of the plant biopolyester cutin as studied by AFM and solid-state NMR. Biophysical Journal 79, 2761–2767. , , , , & (
- 1945) Quantitative saccharification of wood and cellulose. Industrial and Engineering Chemistry – Analytical Edition 17, 35–37. , & (
- 2006) The cutin biopolymer matrix. In Biology of the Plant Cuticle (eds M. Riederer & C. Müller), pp. 126–144. Blackwell Publishing Ltd., Oxford, UK. & (
- 1986) The Botany of Mangroves. Cambridge University Press, Cambridge, UK. (
- 2008) Microwave heating of tea residue yields polysaccharides, polyphenols, and plant biopolyester. Journal of Agricultural and Food Chemistry 56, 11293–11299. , , & (
- 1976) Chemistry of waxes of higher plants. In Chemistry and Biochemistry of Natural Waxes (ed. P.E. Kolattukudy), pp. 236–279. American Elsevier Publishing Co. Inc., New York. (
- 1999) Characterization and biosynthesis of non-degradable polymers in plant cuticles. Planta 208, 181–187. , , & (
- 2000) Monitoring biopolymers present in plant cuticles by FT-IR spectroscopy. Journal of Plant Physiology 156, 419–422. , & (
- 1994) Soil salinity and pH in Japanese mangrove forests and growth of cultivated mangrove plants in different soil conditions. Journal of Plant Research 107, 39–46. , & (
- 1972) Determination of the structure of cutin monomers by a novel depolymerization procedure and combined gas chromatography and mass spectrometry. Biochemistry 11, 1885–1897. & (
- 1998) Biomechanics of isolated plant cuticles. Bot Acta 111, 28–34. & (
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Table S1. Mass spectral data for identification of reduced cutin monomers.
Figure S1. Typical total ion chromatogram (TIC) of reduced cutin monomers obtained by hydrogenolysis of cutin in leaves of S. alba. The TIC from adaxial CM of L4 leaf is shown as representative. The peak numbers were cited as listed in Supporting Information Table S1.
Figure S2. Mass spectra of the TMS ethers of the hexadecane-3-ols (cited as Peak 5 in Supporting Information Table S1 and Supporting Information Fig. S1) derived from cutin hydrogenolysate (bottom) and deuteriolysate (top). Embedded schemes show diagnostic fragment ions produced by α-cleavages. This fragmentation pattern showed that the corresponding cutin monomer was 9(10),16-dihydroxyhexadecanoic acid.
Figure S3. Mass spectra of the TMS ethers of the octadecane-3-ols (cited as Peak 9 in Supporting Information Table S1 and Supporting Information Fig. S1) derived from hydrogenolysate (bottom) and deuteriolysate (top). Embedded schemes show diagnostic fragment ions produced by α-cleavages. This fragmentation pattern showed that the corresponding cutin monomer was 9,10-epoxy-18-hydroxyoctadecanoic acid.
Appendix S1. Mass identification procedure.
|PCE_2482_sm_Data.doc||245K||Supporting info item|
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