Next Article in Journal
The Preparation of Golgi Apparatus-Targeted Polymer Dots Encapsulated with Carbon Nanodots of Bright Near-Infrared Fluorescence for Long-Term Bioimaging
Previous Article in Journal
Multifunctional Spirogyra-hyalina-Mediated Barium Oxide Nanoparticles (BaONPs): Synthesis and Applications
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Molecules
Volume 28
Issue 17
10.3390/molecules28176365
molecules-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Cuticular Waxes and Cutin in Terminalia catappa Leaves from the Equatorial São Tomé and Príncipe Islands

by
Helena Pereira
*,
Rita Simões
and
Isabel Miranda
Centro de Estudos Florestais (CEF), Laboratório Associado Terra, Instituto Superior de Agronomia, Universidade de Lisboa, Tapada da Ajuda, 1349-017 Lisboa, Portugal
*
Author to whom correspondence should be addressed.
Molecules 2023, 28(17), 6365; https://doi.org/10.3390/molecules28176365
Submission received: 7 August 2023 / Revised: 23 August 2023 / Accepted: 28 August 2023 / Published: 31 August 2023
(This article belongs to the Section Chemical Biology)
Browse Figure
Versions Notes

Abstract

:
This study presents for the first time an analysis of the content and chemical composition of the cuticular waxes and cutin in the leaves of the widespread and important tropical species Terminalia catappa. The leaves were collected in the equatorial Atlantic islands of São Tomé and Príncipe, in the Gulf of Guinea. The epicuticular and intracuticular waxes were determined via dichloromethane extraction and their chemical composition via GC-MS analysis, and the content and monomeric composition of cutin were determined after depolymerization via methanolysis. The leaves contained an epidermal cuticular coverage of 52.8 μg cm−2 of the cuticular waxes (1.4% of mass) and 63.3 μg cm−2 (1.5% of mass) of cutin. Cuticular waxes include mainly n-alkanols and fatty acids, with a substantial proportion of terpenes in the more easily solubilized fraction, and sterols in the more embedded waxes. Cutin is mostly constituted by C16 fatty acids and dihydroxyacids, also including aromatic monomers, suggesting a largely linear macromolecular arrangement. The high proportion of triacontanol, α-amyrin, β-amyrin, germanicol, and lupeol in the easily solubilized cuticular fraction may explain the bioactive properties attributed to the T. catappa leaves via the popular medicine, which allows us to consider them as a potential source for the extraction of these compounds.

Graphical Abstract

1. Introduction

Terminalia catappa L. is a tropical tree species in the family Combretaceae that is native to the tropical regions of Asia and northern Australia but was widely spread by humans, and at present, is common in the tropical parts of the Americas and in Africa. It is commonly called the tropical almond or Indian almond, but the names sea almond, wild almond, and umbrella tree are also used. T. catappa is a resilient species used as ornamentals due to its large and striking leaves and graceful canopy, its protection against strong winds, and also for providing nuts for consumption and being a source for local medicine [1,2].
T. catappa L. is a medium to large deciduous tree that grows to about 25 m or above, with an upright, symmetrical crown and long horizontal branches, and a bole that can go up to 150 cm in diameter, often with buttresses up to 3 m in height [3,4]. It is a salt-tolerant species that is very commonly found in coastal and beach areas or on tidal riverbanks in many tropical regions. The tree has been widely planted as an ornamental and shade tree throughout the tropics, often becoming naturalized, but it also grows in a variety of environments, from humid tropical climates to dry conditions.
The leaves are commonly 15–25 cm long and 10–14 cm broad, simple, and petiolate, with a glossy dark green colour that turns pinkish-reddish or yellow-brown before shedding. The leaves' anatomical features have been reported as follows: stomata are absent on the upper surface and present on the lower surface; simple unicellular trichomes are on the lower surface; the upper epidermal cells are barrel-shaped and single layered; the underlying palisade mesophyll is one layered with cylindrical elongated and closely arranged cells with many chloroplasts; and the spongy mesophyll cells are irregularly shaped, with 3–4 layers of thin-walled parenchyma cells with intercellular spaces [5,6,7].
T. catappa is widely known as a source of medicinal products with pharmacological benefits for the treatment of various ailments, mostly using the leaves and bark, while the nutritional value and health benefits of the fruit and seed are also recognized [8,9,10]. The species has become an integral part of traditional medicine within its native range and has been adopted by local communities in the various expansion areas for their application against diverse ailments such as fever, dysentery, asthma, and rheumatisms, to name a few. T. catappa leaves have been researched regarding their chemical composition including flavonoids, tannins, alkaloids, steroids, and carbohydrates [11,12]. Numerous research works also addressed T. catappa extracts’ bioactivity, e.g., its antioxidant, antitoxic, antimicrobial, and antifungal properties [13,14,15,16,17]. A curious application of the Indian almond leaves is by aquarists who keep fish native to tannin-rich waters by placing them in aquaria to mimic the natural conditions of the species (http://www.indianalmondleaves.com/, accessed on 20 June 2023).
No reports have been published on the leaf cuticle of T. catappa, nor on the chemical characterization of its cuticular waxes and cutin. The leaf cuticle is an important boundary membrane established between the plant and the environment, as it is involved in the regulation of non-stomatal water loss and gas exchange, and also confers resistance to several biotic and abiotic stresses [18,19]. The cuticle is a layer covering the epidermal cells that consists of an insoluble polyester matrix named cutin, and cuticular waxes that are soluble in low-polarity solvents [20,21]. Cuticular waxes are composed of two types: the intracuticular waxes embedded in the cutin matrix, and the epicuticular waxes deposited above the cutin as an external layer. Cuticle thickness, proportion of cutin and cuticular waxes, and their chemical compositions show a high degree of variability between the leaves of different species while also playing a role in the interactions between the leaf’s surface and environment, and therefore, at the forefront of the potential responses to the adverse or changing conditions [22,23,24].
This study presents, for the first time, an analysis of the content and chemical composition of the cuticular waxes and cutin in the leaves of Terminalia catappa. The trees were collected in the equatorial Atlantic islands of São Tomé and Príncipe, in the Gulf of Guinea, at three locations. A methodology was developed to determine the proportion of epicuticular and intracuticular waxes and of their chemical composition, as determined via GC-MS analysis, as well as the content and monomeric composition of cutin after depolymerization via methanolysis. It is the aim to enlarge knowledge on the chemical diversity of leaf cuticles, and, specifically for T. catappa, to have a first insight into the leaf cuticle composition of this widespread and important tropical species.

2. Results and Discussion

2.1. Cuticular Waxes Content

The content in the cuticular waxes in T. catappa leaves was determined via dichloromethane (DCM) solubilization via immersion at ambient temperature with a short period to extract the most accessible waxes, here referred to as epicuticular waxes, and a longer period at a higher temperature to further solubilize the waxes, here named as intracuticular waxes. The terms should therefore be taken with caution, and a more accurate designation would refer to the DCM extracts of whole leaves under the given experimental conditions. However, for simplicity, and following the usual terminology in cuticle studies, the terms epicuticular and intracuticular waxes are used here. It is known that the DCM applied on whole leaves does not extract compounds from the internal leaf tissues [25].
The content of the epicuticular and intracuticular waxes in T. catappa adult leaves is given in Table 1. On average, cuticular waxes represent 1.4% of the dry leaves’ mass (the proportion of the epicuticular and intracuticular waxes was 52.9% and 47.1%, respectively). The between-site variation in the content of the cuticular waxes was high in relation to the epicuticular waxes (e.g., 75% coefficient of variation of the mean), while the intracuticular waxes’ content showed less differences.
Regarding leaf coverage, i.e., the amount per unit surface area of the two sides of the leaves, cuticular waxes corresponded to 52.8 μg/cm2. There is a large range of values for the leaf coverage and ratio of epicuticular to intracuticular waxes in different species, with reports of much higher values than those found here for T. catappa or, on the contrary, to much lower values. For instance, Quercus suber leaves have a substantial cuticular wax layer of 154.3–235.1 μg cm−2 [21], Quercus petraea leaves have 101.5–134.5 μg cm−2 and Fagus sylvatica 30.7–55.2 μg cm−2 [26], Quercus ilex leaves 71 μg cm−2 [27], Quercus robur 59 μg cm−2 [28], and Quercus polymorpha 199.4 μg cm−2 [29]. The ratio of epicuticular to intracuticular waxes, that was 1.1 in T. catappa, also varies largely between species with the predominance either of epicuticular waxes or of intracuticular waxes. For instance, the ratio ranged from high ratio values of 6.9 (Clusia flava) to 4.6 (Garcinia spicata), 3.5 (Schefflera elegantissima), and 2.7 (Citrus aurantium) [23].

2.2. Cutin Content

The cutin content of T. catappa adult leaves is given in Table 1. On average, cutin amounted to 1.5% of the dry leaves, corresponding to a leaf coverage of 63.3 μg/cm2. The between-site variation in cutin content was small.
Cutin content in T. catappa leaves is much lower than the 7.1% reported for Quercus suber leaves [30]. It is known that cutin content in leaves varies widely among plant species, and a large range of cutin leaf coverage values has been reported, e.g., 340.6 μg cm−2 in Rhazya stricta [31], 203.8 μg cm−2 in Gossypium barbadense and 170.7 μg cm−2 in Gossypium hirsutum [32], 33.2 μg cm−2 in Hedera helix and 41.5 μg cm−2 in Prunus laurocerasus [33], and between 3 and 5 μg cm−2 for different Camelia spp. [34].

2.3. Composition of Cuticular Waxes

The composition of the epicuticular and intracuticular waxes is given in Table 2 by chemical family and is detailed in Table 3 by compound. It is clear that there is a difference in chemical composition between epicuticular and intracuticular waxes regarding the proportion of long chain lipids and the relative amounts of specific chemical classes and compounds.
Epicuticular waxes have a striking content of terpenes (51.9%, mainly α-amyrin, β-amyrin and germanicol), with n-alkanols (mainly triacontanol) also representing 17.3%, while fatty acids only amount to 5.7%, hydroxyacids 2.4%, and diacids practically absent. Intracuticular waxes are dominated by n-alkanols (39.0%, also with triacontanol as the main compound) and also have a large proportion of sterols (25.2%), while terpenes correspond to 10.5% and fatty acids to 8.5%.
Cuticular waxes include compounds from different chemical families, namely long-chain aliphatics such as fatty acids, aldehydes, primary and secondary alcohols, ketones, alkanes, and cyclic compounds, such as triterpenoids, tocopherols and phytosterols, and aromatic compounds [19,35,36,37]. There is a large diversity in the composition of cuticular waxes of different species. For instance, in Quercus suber leaves, the cuticular wax layer was composed predominantly of triterpenes and aliphatic compounds with 61–72% and 17–23%, respectively [21], while in Quercus ilex, the most abundant components were n-alkanoic acids (38%) and n-alkanols (43–54%), with small amounts of the triterpenoids α-amyrin and β-amyrin [38]; in Quercus robur, the dominating classes were alcohols (about 70%), fatty acids (20%), aldehydes (28%), and several triperpenoids (8%) [39]. In Castanea sativa, the cuticular waxes consisted of a homologous series of wax lipids (esters, aldehydes, primary alcohols, and fatty acids) and large amounts of triterpenoids (α- and β-amyrin and lupeol, while Fagus sylvatica contained only wax lipids, without any triterpenoids [40]. In Eucalyptus camaldulensis and E. globulus, the cyclic compounds constituted about 39% and 76%, respectively [41,42].
A comparison between the chemical composition of epicuticular and intracuticular waxes in T. catappa leaves (Table 2) shows that the epicuticular layer predominantly concentrated triterpenoids and alkanols which were the main long-chain aliphatic compounds, while the intracuticular layer accumulated more alkanols and fatty acids, along with a high content in sterols. This is contrary to reports from other species for which long-chain aliphatic compounds were predominantly concentrated in the epicuticular layer, while triterpenoids were found in the intracuticular layer [43,44]. Similar results were obtained for a short time DCM extraction of Quercus suber leaves for which 95.5% of the extract was composed of n-alkanols and longer extractions were needed to solubilize the terpenes [25].
The substantial presence of triterpenoids in the easily solubilized surface waxes of T. catappa leaves is in line with their application in traditional medicine. In fact, most of the compounds present in the cuticular waxes of T. catappa (Table 3) have biological activities, which may explain the beneficial medicinal use of the leaves’ extracts. Triacontanol, a C30 n-alkanol, is a plant growth regulator found in epicuticular waxes that is involved in overcoming the negative effects of salt stress [45] and that is known as a growth promotor when exogenously applied to a number of plants [46,47]. α- and β-amyrins are triterpenoids (only differing in the placement of the methyl group) that have pharmacological activities exhibiting anti-inflammatory, antidiabetic, and anticancer effects, and are used as precursors for the biosynthesis of valuable bioactive compounds or as a lead compound for drug development effective in diabetes and atherosclerosis [48,49,50,51,52]. Germanicol is a pentacyclic triterpenoid found in various plants that exhibits selective antiproliferative activity against two human colon cancer cells lines mediated via the induction of apoptosis and the suppression of cell migration [53]. Lupeol is also a pharmacologically active pentacyclic triterpenoid with several potential medicinal properties, like anticancer and anti-inflammatory activity.
The high proportion of triacontanol, α-amyrin, β-amyrin, germanicol, and lupeol in the easily solubilized fraction of cuticular waxes may explain the bioactive properties attributed to T. catappa leaves by the popular medicine. It also allows us to consider the leaves as a potential source for these compounds by their targeted extraction.

2.4. Composition of Cutin

The composition of cutin is given in Table 4 by chemical family and is detailed in Table 5 by compound. Cutin composition is dominated by fatty acids (64.8% of the total compounds), mainly including hexadecanoic acid (48.8%), with a substantial amount of hydroxyacids (18.0%), mostly of 10,16-dihydroxyhexadecanoic acid (15.1%), and the presence of aromatics (4.8%), mainly of methyl p-coumarate (2.5%).
This composition is in line with the overall cutin composition, i.e., cutin being a glyceridic polyester of C16 and C18 hydroxyacids and alkanoic acids with bonding to phenolic acids, namely to p-coumaric acid [54,55,56]. The relative proportion of the monomeric classes varies among species. For instance, in Quercus suber, the cutin contains mostly ω-hydroxyacids 44.4%, fatty acids 20.7%, α,ω-diacids 6.5%, and aromatics 12.8% [57], while ω-hydroxyacids represented 78% of the cutin in Prunus laurocerasus, 70% in Citrus aurantium [58], and 34% in Camelina sativa [34]. The proportion of phenolics also varies, e.g., 27% and 16% for Gossypium barbadense and G. hirsutum, respectively [32], and 13% to 21% in Camelina spp. [34].
Cutin provides strength and rigidity to the epidermal layer, and its macromolecular structure is determined via its monomeric composition and the functional groups that allow the establishment of ester bonds, i.e., the carboxylic and hydroxyl groups. In ω-hydroxy acids, the majority of the terminal hydroxyl groups participate in the ester bonds, but only half of the secondary hydroxyl groups are esterified while the number of unesterified carboxyl groups is very small [20,54,59]. The ω-hydroxy acids, with only a single hydroxyl group at the chain’s end, can only contribute to the formation of linear chains, while the presence of mid-chain hydroxyl groups allows the formation of dendritic structures [54].
In the case of T. catappa (Table 5), the cutin is of a C16 type, and the high content of unsubstituted alkanoic acids in comparison with the mid-chain hydroxylated ω-hydroxy acids indicates a predominantly linear structure.

2.5. Experimental Considerations

The determination of cuticular waxes was made using whole leaves via solubilization in dichloromethane, with care to avoid contamination by leaf internal lipids that may be extracted via solvent entering the leaves through surface lesions or cut areas created during leaf preparation. Due to the large size of the T. catappa leaves, an adaptation was made to the solubilization methodology that we adopted previously for Quercus suber leaves [21,60]. A large and shallow glass container was used where the leaves could be immersed flat, and solubilization was made first to the more easily accessible compounds at the leaf surface (here called epicuticular waxes) via a short 5 min immersion at ambient temperature, followed by a 3 h long immersion at a temperature slightly below the boiling point of the solvent (by 3–4 °C) that solubilized the more entrapped material (here called intracuticular waxes). However, the classification as epicuticular and intracuticular waxes, as used here, should be taken with caution since the results refer to the DCM solubles, respectively, by short-ambient and long-hot solubilization conditions.
Dichloromethane was used as solvent, as previously tested, avoiding more polar solvents such as chloroform or acetone that may have an enhanced entrance to the inner leaf tissues [25].
The subsequent determination of cutin requires a reactive step for its depolymerization and monomer solubilization that was made via methanolysis using the previously tested conditions [30]. Since small leaf pieces were required to fit into the reaction vessel, a previous exhaustive Soxhlet extraction with DCM was made to remove all soluble lipid components that could contaminate the cutin compositional analysis. The yield obtained for these leaf internal DCM solubles was 0.6% (Table 1) that contained (Table 6 and Table 7) mostly fatty acids (51.9%, mainly hexadecanoic acid), aromatics (16.0%, with methyl-coumarate and several phenolics), sterols (6.0%, mainly β-sitosterol), alkanols (4.5%), and sugars (3.9%).

3. Materials and Methods

3.1. Sampling

The leaves were collected from mature Terminalia catappa L. trees growing in three locations in the São Tomé and Principe islands: site 1, in São Tomé’s urban coast (0°12′58″ N, 6°35′40″ E); site 2, in the sand front of the islet Ilhéu das Rolas (0°0′1″ N, 6°31′18″ E); and site 3, in the sand beach Boi of Príncipe island (1°40′51″ N, 7°27′36″ E). The islands have a tropical climate with an annual average rainfall of 900 mm with one long rainy season from September to May, and a mean temperature between 22 °C and 26 °C.
The adult leaves were collected from several branches around the canopy below a height of approximately 2 m, from three trees in each site. The sampling was carried out in April 2023, and only complete and undamaged green leaves were taken. Three leaves for each site were chemically analyzed, and the results should therefore be taken as exploratory given this limited sampling. Leaf area was measured for the leaves that were chemically analyzed. The detailed methodology based on the image analysis of the leaves was reported in [21].

3.2. Determination of Cuticular Waxes

The solvent used for the extraction of the cuticular waxes was dichloromethane (DCM) [25]. Only whole leaves without any fracture or damage to the blade were taken. Since the leaves are large and did not fit into the usual chemical lab vessels, a methodological adaptation was made to our previous experimental procedures by using a shallow rectangular glass vessel (30 cm × 20 cm × 5 cm) where the leaves were immersed flat in the solvent. The leaves were first carefully dried at 60 °C overnight and weighted.
The epicuticular waxes were solubilized via a short 5 min immersion at ambient temperature (23 °C) in 500 mL of solvent. The solvent was taken, evaporated in a rotavapor, and kept for chemical analysis. The extracted leaves were dried at 60 °C overnight and weighted.
The intracuticular waxes were extracted from the same leaves with 500 mL of dichloromethane using the same glass vessel in a water bath at 36 °C during 3 h. The solvent was taken, concentrated, and kept for chemical analysis. The leaves were dried at 60 °C overnight.
With the adopted experimental procedure, the cuticular waxes were extracted from both surfaces of the leaves. The amount of solubilized cuticular material was determined and expressed on a dry weight basis (g/1000 g dry leaf mass) and on a leaf surface area (as μg/cm2) corresponding to a leaf surface coverage, with the area being the two-sided leaf surface area. The detailed methodology was already reported, showing that little amounts of internal lipids or polysaccharides were solubilized under these conditions [25].

3.3. Determination of Cutin Content

In order to perform the depolymerization of cutin via methanolysis, it was necessary to cut the leaves into smaller pieces that would fit into the reaction vessel. This was made via the manual shredding of the leaves into about 1 cm2 pieces. However, in order to avoid any contamination in the chemical analysis of cutin from the internal lipids that would be accessed and solubilized via the reaction solvent, a previous Soxhlet extraction with dichloromethane was made during 6 h. The solvent was evaporated and kept for analysis, and the leaf pieces were dried at 60 °C. The extracted material obtained in this step is here called leaf internal DCM solubles.
The depolymerization of cutin was carried out via transesterification with a sodium methoxide (NaOMe)-catalyzed methanolysis applied to the extracted leaves. The protocol followed the methodology applied to the cork suberin depolymerization [61] that was previously adapted to the cutin determination in cork oak leaves [57].
A sample of approximately 1.5 g extracted leaves was refluxed during 3 h with 100 mL of a methanolic 3% NaOCH3 solution, filtrated, washed with CH3OH, refluxed again with 100 mL of CH3OH during 15 min, filtrated, and then the combined filtrates were acidified to pH 6 with 2 M H2SO4 and evaporated to dryness. The residue was suspended in 50 mL of water and extracted three times with dichloromethane (50 mL each). The combined dichloromethane extracts were dried over anhydrous Na2SO4 and evaporated to dryness.
Cutin was quantified by determining the mass loss via methanolysis after drying and weighing the leaves’ residue, and it was expressed in percentage form of the initial dry weight basis (g/1000 g dry leaf mass) and on a leaf surface area (as μg/cm2), corresponding to a leaf surface coverage, with the area being the two-sided leaf surface area.

3.4. Chemical Composition of Cuticular Extracts and of Cutin

The extracts of the epicuticular and intracuticular waxes, the leaf internal DCM solubles, and the cutin monomers obtained via transesterification of the extracted leaves were solubilized in dichloromethane and derivatized into silylated derivatives for GC-MS analysis.
Aliquots (5 mL) of the dichloromethane extracts were evaporated under N2 flow and dried overnight at ambient conditions under a vacuum. The samples were derivatized in 100 μL of pyridine with 100 μL of bis(trimethylsilyl) trifluoroacetamide at 60 °C for 30 min, in order to trimethylsilylate (TMS) the hydroxyl and carboxyl groups into ethers and esters, respectively.
The derivatized samples were immediately injected in a GC-MS Agilent 5973 MSD with the following GC conditions: Zebron 7HG-G015-02 column (30 m, 0.25 mm; ID, 0.1 μm film thickness), flow of 1 mL/min, injector of 280 °C, oven temperature program; 100 °C (1 min); a rate of 8 °C/min up to 250 °C; a rate of 5 °C/min up to 300 °C (5 min); a rate of 5 °C/min up to 350 °C (5 min); and a rate of 10 °C/min up to 380 °C (5 min). The MS source was kept at 220 °C and the electron impact mass spectra (EIMS) was taken at 70 eV. The compounds were identified as TMS derivatives by comparing with a GC-MS spectral library (Wiley, NIST, Gaithersburg, MD, USA) and with the published data, as reported in [21]. A semi-quantitative approach to the chemical composition was taken by using the integration of the peak area in the GC-MS total ion chromatograms, where each peak was quantified in the area proportion of the total chromatogram area. Duplicate analyses were made (with the standard deviation below 5%).

4. Conclusions

The widespread tropical species Terminalia catappa has an epidermal cuticular coverage of 52.8 μg cm−2 of cuticular waxes and 63.3 μg cm−2 of cutin. Cuticular waxes include mainly n-alkanols and fatty acids, with a substantial proportion of terpenes in the more easily solubilized fraction, and sterols in the more embedded waxes.
Cutin is mostly constituted by C16 fatty acids and dihydroxyacids, also including aromatic monomers, suggesting a largely linear macromolecular arrangement.
The high proportion of compounds with bioactive properties such as triacontanol, α-amyrin, β-amyrin, germanicol, and lupeol in the easily solubilized cuticular fraction allows us to consider the T. catappa leaves as a potential source for their extraction.

Author Contributions

Conceptualization, H.P.; methodology, I.M. and R.S.; validation, H.P., I.M., and R.S.; formal analysis, R.S.; investigation, R.S.; data curation, R.S., H.P., and I.M.; writing—original draft preparation, H.P., I.M., and R.S.; writing—review and editing, H.P. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Portuguese Foundation for Science and Technology (FCT) through the funding of the Forest Research Centre (UIDB/00239/2020). Rita Simões acknowledges a doctoral scholarship from FCT with the SUSFOR Doctoral Programme (PD/BD/128259/2016).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original research data are available from the authors.

Acknowledgments

We thank Joaquina Silva for her technical assistance in the chemical analysis.

Conflicts of Interest

The authors declare no conflict of interest.

Sample Availability

Not applicable.

References

  1. Anand, A.V.; Divya, N.; Kotti, P.P. An updated review of Terminalia catappa. Pharmacogn. Rev. 2015, 9, 93–98. [Google Scholar] [CrossRef] [PubMed]
  2. Yada, V.; Munin, A. Phytochemical characteristic and physical analysis of some medicinal plants. J. Physiol. 2011, 3, 10–14. Available online: http://journal-phytology.com/ (accessed on 20 June 2023).
  3. Marjenah, M.; Putri, N.P. Morphological characteristics and physical environment of Terminalia catappa in East Kalimantan, Indonesia. Asian J. For. 2017, 1, 33–39. [Google Scholar] [CrossRef]
  4. Thomson, L.A.J.; Evans, B. Terminalia catappa. In Species Profiles for Pacific Island Agroforestry: Permanent Agriculture Resources; Elevitch, C.R., Ed.; Agroforestry: Hōlualoa, HI, USA, 2006; Available online: http://www.traditionaltree.org/ (accessed on 20 June 2023).
  5. Ekeke, C.; Agbagwa, I.O. Epidermal structures and stomatal ontogeny in Terminalia catappa L. (Combretaceae). Int. J. Bot. 2015, 11, 1–9. [Google Scholar] [CrossRef]
  6. Punwong, P.; Juprasong, Y.; Traiperm, P. Effects of an oil spill on the leaf anatomical characteristics of a beach plant (Terminalia catappa L.). Environ. Sci. Pollut. Res. 2017, 24, 21821–21828. [Google Scholar] [CrossRef]
  7. Zan, B. Study on morphology, anatomy and phytochemical test of Terminalia catappa L. leaves. 3rd Myanmar Korea Conf. Res. J. 2020, 3, 67. [Google Scholar]
  8. Oliveira, J.T.A.; Vasconcelos, I.M.; Bezerra, L.C.N.M.; Silveira, S.B.; Monteiro, A.C.O.; Moreira, R.A. Composition and nutritional properties of seeds from Pachira aquatica Aubl, Sterculia striata StHil et Naud and Terminalia catappa Linn. Food Chem. 2000, 70, 185–191. [Google Scholar] [CrossRef]
  9. Nagappa, A.N.; Thakurdesai, P.A.; Venkat Rao, N.; Singh, J. Antidiabetic effect of Terminalia catappa Linn fruits. Ethnopharmacology 2003, 88, 45–50. [Google Scholar] [CrossRef]
  10. Ramachandran, S.; Singh, S.K.; Larroche, C.; Soccol, C.R.; Pandey, A. Oil cakes and their biotechnological applications—A review. Bioresour. Technol. 2007, 98, 2000–2009. [Google Scholar] [CrossRef]
  11. Ko, T.F.; Weng, Y.M.; Chiou, Y.Y. Squalene content and antioxidant activity of Terminalia catappa leaves and seeds. J. Agric. Food Chem. 2002, 50, 5343–5348. [Google Scholar] [CrossRef]
  12. Tanaka, T.; Nonaka, G.I.; Nishioka, I. Tannins and related compounds. Isolation and characterization of four new hydrolyzable tannins, terflavins A and B, tergallagin and tercatain from the leaves of Terminalia catappa L. Chem. Pharm. Bull. 1986, 34, 1039–1049. [Google Scholar] [CrossRef]
  13. Chyau, C.C.; Ko, P.T.; Mau, J.L. Antioxidant properties of aqueous extracts from Terminalia catappa leaves. Lebensm. Wiss. Technol. 2006, 39, 1099–1108. [Google Scholar] [CrossRef]
  14. Fan, Y.M.; Xu, L.Z.; Gao, J.; Wang, Y.; Tang, X.H.; Zhao, X.N.; Zhang, Z.X. Phytochemical and antiinflamatory studies on Terminalia catappa. Fitoterapia 2004, 75, 253–260. [Google Scholar] [CrossRef]
  15. Mandloi, S.; Mishra, R.; Varma, R.; Varughese, B.; Tripathi, J. A study on phytochemical and antifungal activity of leaf extracts of Terminalia catappa. Int. J. Pharm. Bio Sci. 2013, 4, 1385–1393. [Google Scholar]
  16. Nair, R.; Chanda, S. Antimicrobial activity of Terminalia catappa, Manilkarazapota and Piper betel leaf extract. Indian J. Pharm. Sci. 2008, 70, 390–393. [Google Scholar] [CrossRef]
  17. Terças, A.G.; Monteiro, A.S.; Moffa, E.B.; Santos, J.R.A.; Sousa, E.M.; Pinto, A.R.B.; Costa, P.C.S.; Borges, A.C.R.; Torres, L.M.B.; Barros Filho, A.K.D.; et al. Phytochemical characterization of Terminalia catappa Linn. extracts and their antifungal activities against Candida spp. Front. Microbiol. 2017, 8, 595. [Google Scholar] [CrossRef] [PubMed]
  18. Buschhaus, C.; Jetter, R. Composition differences between epicuticular and intracuticular wax substructures: How do plants seal their epidermal surfaces? J. Exp. Bot. 2011, 62, 841–853. [Google Scholar] [CrossRef] [PubMed]
  19. Yeats, T.H.; Rose, J.K. The formation and function of plant cuticles. Plant Physiol. 2013, 163, 5–20. [Google Scholar] [CrossRef]
  20. Pollard, M.; Beisson, F.; Li, Y.; Ohlrogge, J.B. Building lipid barriers: Biosynthesis of cutin and suberin. Trends Plant Sci. 2008, 13, 236–246. [Google Scholar] [CrossRef]
  21. Simões, R.; Rodrigues, A.; Ferreira-Dias, S.; Miranda, I.; Pereira, H. Chemical composition of cuticular waxes and pigments and morphology of leaves of Quercus suber trees of different provenance. Plants 2020, 9, 1165. [Google Scholar] [CrossRef]
  22. Bourgault, R.; Matschi, S.; Vasquez, M.; Qiao, P.; Sonntag, A.; Charlebois, C.; Mohammadi, M.; Scanlon, M.J.; Smith, L.G.; Molina, I. Constructing functional cuticles: Analysis of relationships between cuticle lipid composition, ultrastructure and water barrier function in developing adult maize leaves. Ann. Bot. 2020, 125, 79–91. [Google Scholar] [CrossRef] [PubMed]
  23. Jetter, R.; Riederer, M. Localization of the transpiration barrier in the epi- and intracuticular waxes of eight plant species: Water transport resistances are associated with fatty acyl rather than alicyclic components. Plant Physiol. 2016, 170, 921–993. [Google Scholar] [CrossRef]
  24. Tomasi, P.; Dyer, J.M.; Jenks, M.A.; Abdel-Haleem, H. Characterization of leaf cuticular wax classes and constituents in a spring Camelina sativa diversity panel. Ind. Crops Prod. 2018, 112, 247–251. [Google Scholar] [CrossRef]
  25. Simões, R.; Miranda, I.; Pereira, H. The influence of solvent and extraction time on yield and chemical selectivity of cuticular waxes from Quercus suber leaves. Processes 2022, 10, 2270. [Google Scholar] [CrossRef]
  26. Bahamonde, H.A.; Gil, L.; Fernández, V. Surface properties and permeability to calcium chloride of Fagus sylvatica and Quercus petraea leaves of different canopy heights. Front. Plant Sci. 2018, 9, 494. [Google Scholar] [CrossRef] [PubMed]
  27. Martins, C.M.C.; Mesquita, S.M.M.; Vaz, W.L.C. Cuticular waxes of the holm (Quercus ilex L. subsp ballota (Desf.) Samp.) and cork (Q. suber L.) oaks. Phytochem. Anal. 1999, 10, 1–5. [Google Scholar]
  28. Prasad, R.B.N.; Gülz, P.G. Surface structure and chemical composition of leaf waxes from Quercus robur L., Acer pseudoplatanus L. and JugIans regia L. Z. Naturforschung C 1990, 45, 813–817. [Google Scholar] [CrossRef]
  29. Maiti, R.; Rodriguez, H.G.; Sarkar, N.C.; Kumari, A. Biodiversity in leaf chemistry (pigments, epicuticular wax and leaf nutrients) in woody plant species in north-eastern Mexico: A synthesis. Forest Res. 2016, 5, 170. [Google Scholar] [CrossRef]
  30. Simões, R.; Miranda, I.; Pereira, H. Variation in leaf cutin content and chemical composition along one annual cycle in the Mediterranean cork oak (Quercus suber L.). Forests 2023, 14, 334. [Google Scholar] [CrossRef]
  31. Schuster, A.-C.; Burghardt, M.; Alfarhan, A.; Bueno, A.; Hedrich, R.; Leide, J.; Thomas, J.; Riederer, M. Effectiveness of cuticular transpiration barriers in a desert plant at controlling water loss at high temperatures. AoB Plants 2016, 8, plw027. [Google Scholar] [CrossRef]
  32. Tomasi, P.; Herritt, M.T.; Jenks, M.A.; Thompson, A.L. Quantification of leaf wax and cutin monomer composition in Pima (Gossypium barbadense L.) and upland (G hirsutum L.) cotton. Ind. Crops Prod. 2021, 169, 113670. [Google Scholar] [CrossRef]
  33. Graça, J.; Schreiber, L.; Rodrigues, J.; Pereira, H. Glycerol and glyceryl esters of omega-hydroxyacids in cutins. Phytochemistry 2002, 61, 205–215. [Google Scholar] [CrossRef]
  34. Tomasi, P.; Wang, H.; Lohrey, G.T.; Park, S.; Dyer, J.M.; Jenks, M.A.; Abdel-Haleem, H. Characterization of leaf cuticular waxes and cutin monomers of Camelina sativa and closely-related Camelina species. Ind. Crops Prod. 2017, 98, 130–138. [Google Scholar] [CrossRef]
  35. Domínguez, E.; Heredia-Guerrero, J.A.; Heredia, A. The biophysical design of plant cuticles: An overview. New Phytol. 2011, 189, 938–949. [Google Scholar] [CrossRef] [PubMed]
  36. Jetter, R.; Kunst, L.; Samuels, A.L. Composition of plant cuticular waxes. In Biology of the Plant Cuticle; Riederer, M., Müller, C., Eds.; Blackwell Publishing Ltd.: Oxford, UK, 2006; pp. 145–181. [Google Scholar] [CrossRef]
  37. Sharma, P.; Kothari, S.L.; Rathore, M.S.; Gour, V.S. Properties variations roles potential applications of epicuticular wax: A review. Turk. J. Bot. 2018, 42, 135–149. [Google Scholar] [CrossRef]
  38. Vaz, M.; Moroco, J.; Ribeiro, N.; Gazarini, L.C.; Pereira, J.S.; Chaves, M.M. Leaf-level responses to light in two co-occurring Quercus (Quercus ilex and Quercus suber): Leaf structure, chemical composition and photosynthesis. Agrofor. Syst. 2011, 82, 173–181. [Google Scholar] [CrossRef]
  39. Gülz, P.G.; Müller, E. Seasonal variation in the composition of epicuticular waxes of Quercus robur leaves. Z. Naturforschung C 1992, 47, 800–806. [Google Scholar] [CrossRef]
  40. Gülz, P.G.; Müller, E.; Herrmann, T. Chemical composition and surface structures of epicuticular leaf waxes from Castanea sativa and Aesculus hippocastanum. Z. Naturforschung C 1992, 47, 661–666. [Google Scholar] [CrossRef]
  41. Gülz, P.G.; Prasad, R.B.N.; Müller, E. Surface structures and chemical composition of epicuticular waxes during leaf development of Fagus sylvatica L. Z. Naturforschung C 1992, 47, 190–196. [Google Scholar] [CrossRef]
  42. Guzmán, P.; Fernández, V.; Graça, J.; Cabral, V.; Kayali, N.; Khayet, M.; Gil, L. Chemical and structural analysis of Eucalyptus globulus and E. camaldulensis leaf cuticles: A lipidized cell wall region. Front. Plant Sci. 2014, 5, 481. [Google Scholar] [CrossRef]
  43. Guhling, O.; Kinzler, C.; Dreyer, M.; Bringmann, G.; Jetter, R. Surface composition of myrmecophytic plants: Cuticular wax and glandular trichomes on leaves of Macaranga tanarius. J. Chem. Ecol. 2005, 31, 2325–2343. [Google Scholar] [CrossRef]
  44. Jetter, R.; Schäffer, S.; Riederer, M. Leaf cuticular waxes are arranged in chemically and mechanically distinct layers: Evidence from Prunus laurocerasus L. Plant Cell Environ. 2000, 23, 619–623. [Google Scholar] [CrossRef]
  45. Verma, T.; Bhardwaj, S.; Singh, J.; Kapoor, D.; Prasad, R. Triacontanol as a versatile plant growth regulator in overcoming negative effects of salt stress. J. Agric. Food Res. 2022, 10, 100351. [Google Scholar] [CrossRef]
  46. Naeem, M.; Khan, M.M.A.; Moinuddin. Triacontanol: A potent plant growth regulator in agriculture. J. Plant Interact. 2011, 7, 129–142. [Google Scholar] [CrossRef]
  47. Mian, G.; Belfiore, N.; Musetti, R.; Tomasi, D.; Cantone, P.; Lovat, L.; Lupinelli, S.; Iacumin, L.; Celotti, E.; Golinelli, F. Effect of a triacontanol-rich biostimulant on the ripening dynamic and wine must technological parameters in Vitis vinifera cv. ‘Ribolla Gialla’. Plant Phys. Biochem. 2022, 188, 60–69. [Google Scholar] [CrossRef] [PubMed]
  48. Askari, V.R.; Fereydouni, N.; Baradaran Rahimi, V.; Askari, N.; Sahebkar, A.H.; Rahmanian-Devin, P.; Samzadeh-Kermani, A. β-Amyrin, the cannabinoid receptors agonist, abrogates mice brain microglial cells inflammation induced by lipopolysaccharide/interferon-γ and regulates Mφ1/Mφ2 balances. Biomed. Pharmacother. 2018, 101, 438–446. [Google Scholar] [CrossRef]
  49. Neto, S.F.; Prada, A.L.; Achod, L.D.R.; Torquato, H.F.V.; Lima, C.S.; Paredes-Gamero, E.J.; Moraes, M.O.; Lima, E.S.; Sosa, E.H.; de Souza, T.P.; et al. α-Amyrin-loaded nanocapsules produce selective cytotoxic activity in leukemic cells. Biomed. Pharmacother. 2021, 139, 111656. [Google Scholar] [CrossRef]
  50. Santos, F.A.; Frota, J.T.; Arruda, B.R.; de Melo, T.S.; de Carvalho Almeida da Silva, A.A.; de Castro Brito, G.A.; Chaves, M.H.; Rao, V.S. Antihyperglycemic and hypolipidemic effects of α, β-amyrin, a triterpenoid mixture from Protium heptaphyllum in mice. Lipids Health Dis. 2012, 11, 98. [Google Scholar] [CrossRef]
  51. Seki, H.; Ohyama, K.; Sawai, S.; Mizutani, M.; Ohnishi, T.; Sudo, H.; Akashi, T.; Aoki, T.; Saito, K.; Muranaka, T. Licorice β-amyrin 11-oxidase, a cytochrome P450 with a key role in the biosynthesis of the triterpene sweetener glycyrrhizin. Proc. Natl. Acad. Sci. USA 2008, 105, 14204–14209. [Google Scholar] [CrossRef]
  52. Yang, W.; Chen, X.; Li, Y.; Guo, S.; Wang, Z.; Yu, X. Advances in pharmacological activities of terpenoids. Nat. Prod. Commun. 2020, 15, 1–13. [Google Scholar] [CrossRef]
  53. Hu, Y.L.; Wang, X.B.; Chen, D.D.; Guo, X.J.; Yang, Q.J.; Dong, L.H.; Cheng, L. Germanicol induces selective growth inhibitory effects in human colon HCT-116 and HT29 cancer cells through induction of apoptosis, cell cycle arrest and inhibition of cell migration. J. BUON 2016, 21, 626–632. [Google Scholar]
  54. Fich, E.A.; Segerson, N.A.; Rose, J.K. The plant polyester cutin: Biosynthesis, structure, and biological roles. Annu. Rev. Plant Biol. 2016, 67, 207–233. [Google Scholar] [CrossRef]
  55. Li-Beisson, Y.; Verdier, G.; Xu, L.; Beisson, F. Cutin and suberin polyesters. In Essential for Life Science; John Wiley & Sons, Ltd.: Chichester, UK, 2016; pp. 1–12. [Google Scholar] [CrossRef]
  56. Riley, R.G.; Kolattukudy, P.E. Evidence for covalently attached p-voumaric acid and ferulic acid in cutins and suberins. Plant Physiol. 1975, 56, 650–654. [Google Scholar] [CrossRef] [PubMed]
  57. Simões, R.; Miranda, I.; Pereira, H. Chemical composition of leaf cutin in six Quercus suber provenances. Phytochemistry 2021, 181, 112570. [Google Scholar] [CrossRef] [PubMed]
  58. Graça, J.; Lamosa, P. Linear branched poly(ω-hydroxyacid) esters in plant cutins. J. Agric. Food Chem. 2010, 58, 9666–9674. [Google Scholar] [CrossRef]
  59. Kolattukudy, P.E. Biosynthetic pathways of cutin and waxes, and their sensitivity to environmental stresses. In Plant Cuticles: An Integrated Functional Approach; Kerstiens, G., Ed.; BIOS Scientific Publishers: Oxford, UK, 1996; pp. 83–108. [Google Scholar]
  60. Simões, R.; Miranda, I.; Pereira, H. Effect of seasonal variation on leaf cuticular waxes’ composition in the Mediterranean cork oak (Quercus suber L.). Forests 2022, 13, 1236. [Google Scholar] [CrossRef]
  61. Pereira, H. Chemical composition and variability of cork from Quercus suber L. Wood Sci. Technol. 1988, 22, 211–218. [Google Scholar] [CrossRef]
Table 1. Chemical characterization of the leaves of Terminalia catappa of adult trees in three sites in the islands of São Tomé and Príncipe, regarding epicuticular and intracuticular waxes and cutin, in mg/g of dry leaves and in μg/cm2 of leaf surface. The leaf internal dichloromethane (DCM) solubles are also reported. Mean and standard deviation (±std) are also included.
Table 1. Chemical characterization of the leaves of Terminalia catappa of adult trees in three sites in the islands of São Tomé and Príncipe, regarding epicuticular and intracuticular waxes and cutin, in mg/g of dry leaves and in μg/cm2 of leaf surface. The leaf internal dichloromethane (DCM) solubles are also reported. Mean and standard deviation (±std) are also included.
Cuticular Lipid ContentSite 1Site 2Site 3Mean ± Std
Cuticular waxes
  Epicuticular waxes (mg/g)13.34.93.37.2 ± 5.4
  Epicuticular waxes (μg/cm2)36.426.014.225.6 ± 11.1
  Intracuticular waxes (mg/g)5.07.36.86.4 ± 1.0
  Intracuticular waxes (μg/cm2)13.838.229.727.2 ± 12.4
  Total (mg/g)18.312.210.113.6 ± 3.5
  Total (μg/cm2)50.264.343.952.8 ± 10.4
Internal DCM solubles (mg/g)3.86.78.16.2 ± 2.2
Cutin (mg/g)12.415.716.915.0 ± 2.3
Cutin (μg/cm2)33.983.372.763.3 ± 21.2
Table 2. Composition by chemical class of the cuticular waxes (epicuticular and intracuticular) of leaves of Terminalia catappa adult trees in three sites in the islands of São Tomé and Príncipe, as a percentage of the total peak areas in the GC-MS chromatogram. Mean and standard deviation (±std) are also included.
Table 2. Composition by chemical class of the cuticular waxes (epicuticular and intracuticular) of leaves of Terminalia catappa adult trees in three sites in the islands of São Tomé and Príncipe, as a percentage of the total peak areas in the GC-MS chromatogram. Mean and standard deviation (±std) are also included.
Chemical ClassEpicuticular WaxesMean ± StdIntracuticular WaxesMean ± Std
Site 1Site 2Site 3Site 1Site 2Site 3
n-Alkanols23.518.010.317.3 ± 6.654.433.529.039.0 ± 13.6
n-Alkanes2.311.20.14.5 ± 5.92.11.17.1 3.4 ± 3.2
Fatty acids9.45.52.35.7 ± 3.65.310.69.5 8.5 ± 2.8
Diacids2.50.00.00.8 ± 1.40.00.40.00.1 ± 0.2
Hydroxyacids7.30.00.02.4 ± 4.20.10.30.3 0.2 ± 0.1
Aromatic compounds0.71.31.81.3 ± 0.60.30.60.2 0.4 ± 0.2
Sterols0.01.10.70.6 ± 0.4625.224.625.725.2 ± 0.6
Terpenes38.647.769.351.9 ± 15.86.713.011.910.5 ± 3.4
Sugars2.80.20.01.0 ± 1.60.01.80.00.6 ± 1.0
Others0.61.60.91.0 ± 0.41.54.42.22.7 ± 1.5
Total identified88.186.785.386.7 ± 1.495.590.185.886.7 ± 1.4
Table 3. Composition of the cuticular waxes (epicuticular and intracuticular) of leaves of Terminalia catappa adult trees as the averages of three sites in the islands of São Tomé and Príncipe, as a percentage of the total peak areas in the GC-MS chromatogram. Only compounds above 1% of total peak area at least in one site are included. Mean and standard deviation (±std) are also included.
Table 3. Composition of the cuticular waxes (epicuticular and intracuticular) of leaves of Terminalia catappa adult trees as the averages of three sites in the islands of São Tomé and Príncipe, as a percentage of the total peak areas in the GC-MS chromatogram. Only compounds above 1% of total peak area at least in one site are included. Mean and standard deviation (±std) are also included.
CompoundsEpicuticular WaxesMean ± StdIntracuticular WaxesMean ± Std
Site 1Site 2Site 3Site 1Site 2Site 3
Alcohols
Octacosanol0.81.40.30.8 ± 0.41.30.62.11.3 ± 0.6
Triacontanol20.214.39.214.6 ± 4.547.325.221.531.4 ± 13.9
Dotriacontanol1.43.40.31.3 ± 0.95.24.94.34.8 ± 0.4
Tetratriacontanol0.00.00.50.2 ± 0.20.62.71.01.4 ± 0.9
Alkanes
Octacosane0.86.60.02.5 ± 3.00.70.03.41.4 ± 1.5
Triacontane1.54.30.01.9 ± 1.81.31.13.72.1 ± 1.2
Fatty acids
Hexadecanoic acid3.31.11.01.8 ± 1.01.73.53.83.0 ± 0.9
9,12-Octadecadienoic acid0.20.00.00.1 ± 0.10.31.40.50.7 ± 0.5
9-Octadecenoic acid0.10.40.00.2 ± 0.20.31.51.31.0 ± 0.5
Octadecanoic acid1.30.40.50.7 ± 0.40.30.61.00.6 ± 0.3
Dotriacontanoic acid0.01.70.00.6 ± 0.80.81.50.30.9 ± 0.5
α,ω-Diacids
9,10-Dihydroxyoctadecanedioic acid, dimethyl ester1.90.00.00.6 ± 0.90.00.40.00.1 ± 0.2
ω-Hydroxy fatty acids
9,10,18-Trihydroxyoctadecanoic acid, methyl ester1.20.00.00.6 ± 0.9
22-Hidroxydocosanoic, methyl ester5.80.00.01.9 ± 3.30.10.20.30.2 ± 0.1
Steroids
Stigmasterol0.01.10.70.6 ± 0.52.22.03.52.5 ± 0.7
β-sitosterol 22.521.921.221.9 ± 0.5
Terpenes
Phytol0.00.50.10.2 ± 0.20.20.91.40.6 ± 0.5
Squalene 0.21.32.11.2 ± 0.8
β-Amyrone2.40.30.31.0 ± 1.0
β-Amyrin7.97.414.59.9 ± 3.33.42.62.72.7 ± 0.6
Germanicol6.410.40.05.6 ± 4.3
α-Amyrin17.211.231.720.0 ± 8.61.11.61.81.5 ± 0.3
Lupeol4.14.79.46.0 ± 2.4
Betulinol0.35.74.03.3 ± 2.3
Oleanolic acid0.15.72.11.9 ± 1.8
Betulinic acid0.11.70.80.9 ± 0.6
Ursolic acid 0.01.20.20.5 ± 0.5
Corosolic acid 0.43.41.61.8 ± 1.2
Phytyl octadecanoate (C18:0) 0.41.21.81.1 ± 0.6
Sugars
meso-erythritol 0.01.80.00.6 ± 0.6
Ribitol2.80.00.00.9 ± 1.3
Others
1(3H)-Isobenzofuranone, 3-methyl-3-(2,4,6-trimethoxyphenyl)-0.11.10.60.6 ± 0.50.63.21.91.9 ± 1.3
Total identified compounds80.383.784.082.7 ± 2.192.185.781.986.5 ± 5.2
Table 4. Composition by chemical class of cutin of leaves of Terminalia catappa adult trees in three sites in the islands of São Tomé and Príncipe, as percentage of the total peak areas in the GC-MS chromatogram. Mean and standard deviation (±std) are also included.
Table 4. Composition by chemical class of cutin of leaves of Terminalia catappa adult trees in three sites in the islands of São Tomé and Príncipe, as percentage of the total peak areas in the GC-MS chromatogram. Mean and standard deviation (±std) are also included.
Chemical ClassCutinMean ± Std
Site 1Site 2Site 3
n-Alkanols0.41.00.80.7 ± 0.3
Fatty acids72.565.756.364.8 ± 8.1
Diacids1.70.52.01.4 ± 0.8
Hydroxyacids9.516.328.318.0 ± 9.5
Aromatic compounds4.16.53.94.8 ± 1.5
Others2.32.11.92.1 ± 0.2
Total identified compounds90.592.193.291.9 ± 1.4
Table 5. Composition of the cutin of leaves of Terminalia catappa adult trees as the averages of three sites in the islands of São Tomé and Príncipe, as a percentage of the total peak areas in the GC-MS chromatogram. Only compounds above 1% of total peak area at least in one site are represented. Mean and standard deviation (±std) are also included.
Table 5. Composition of the cutin of leaves of Terminalia catappa adult trees as the averages of three sites in the islands of São Tomé and Príncipe, as a percentage of the total peak areas in the GC-MS chromatogram. Only compounds above 1% of total peak area at least in one site are represented. Mean and standard deviation (±std) are also included.
Chemical ClassCutinMean ± Std
Site 1Site 2Site 3
Fatty acids
Hexadecanoic acid4.05.18.86.0 ± 2.5
9-Decenoic acid3.14.92.63.5 ± 1.2
Hexadecanoic acid, methyl ester 57.650.438.548.8 ± 9.6
Octadecanoic acid, methyl ester2.33.23.12.8 ± 0.5
EIcosanoic acid, methyl ester 1.20.51.00.9 ± 0.4
ω-Hydroxy fatty acids
16-Hydroxyhexadecanoic acid1.92.63.92.8 ± 1.0
10,16-Dihydroxyhexadecanoic acid7.413.624.215.1 ± 8.5
Phenolics
Benzoic acid 0.71.00.80.8 ± 0.2
Benzeneacetic acid1.00.00.00.3 ± 0.6
Methyl p-coumarate 0.51.80.91.0 ± 0.7
Methyl p-coumarate (isomer)0.92.51.31.5 ± 0.8
Methyl isoferulate0.01.10.00.4 ± 0.6
Others
2-Pentadecenone, 6,10,14-trimethyl-1.40.00.00.5 ± 0.8
Methyl 14-methoxyhexadecanoate1.02.11.81.6 ± 0.6
Total identified compounds83.088.786.986.2 ± 2.9
Table 6. Composition by chemical class of the leaf internal dichloromethane (DCM) solubles after extraction of cuticular waxes (epicuticular and intracuticular) of leaves of Terminalia catappa adult trees in three sites in the islands of São Tomé and Príncipe, as a percentage of the total peak areas in the GC-MS chromatogram. Mean and standard deviation (±std) are also included.
Table 6. Composition by chemical class of the leaf internal dichloromethane (DCM) solubles after extraction of cuticular waxes (epicuticular and intracuticular) of leaves of Terminalia catappa adult trees in three sites in the islands of São Tomé and Príncipe, as a percentage of the total peak areas in the GC-MS chromatogram. Mean and standard deviation (±std) are also included.
Chemical ClassLeaf Internal DCM SolublesMean ± Std
Site 1Site 2Site 3
n-Alkanols5.22.65.64.5 ± 1.6
n-Alkanes0.80.32.41.2 ± 1.1
Fatty acids53.150.152.451.9 ± 1.6
Diacids1.21.41.61.4 ± 0.2
Aromatic compounds23.216.38.416.0 ± 6.1
Sterols8.23.56.46.0 ± 2.4
Terpenes0.30.01.20.5 ± 0.5
Sugars1.12.97.73.9 ± 2.8
Total identified compounds93.178.383.885.1 ± 6.1
Table 7. Chemical composition of the leaf internal dichloromethane (DCM) solubles after extraction of cuticular waxes (epicuticular and intracuticular) of leaves of Terminalia catappa adult trees in three sites in the islands of São Tomé and Príncipe, as a percentage of the total peak areas in the GC-MS chromatogram. Only compounds above 1% of total peak area at least in one site are included. Mean and standard deviation (±std) are also included.
Table 7. Chemical composition of the leaf internal dichloromethane (DCM) solubles after extraction of cuticular waxes (epicuticular and intracuticular) of leaves of Terminalia catappa adult trees in three sites in the islands of São Tomé and Príncipe, as a percentage of the total peak areas in the GC-MS chromatogram. Only compounds above 1% of total peak area at least in one site are included. Mean and standard deviation (±std) are also included.
CompoundLeaf internal DCM SolublesMean ± Std
Site 1Site 2Site 3
Alchools
1,2-Hexadecanediol1.61.70.91.4 ± 0.4
Dodecanol0.40.01.10.5 ± 0.5
Triacontanol2.40.52.01.6 ± 1.0
Dotriacontanol0.70.31.00.6 ± 0.4
Alkanes
Octacosane0.30.11.10.5 ± 0.5
Triacontane0.50.21.40.7 ± 0.6
Fatty acids
Hexanoic acid1.90.00.00.6 ± 1.1
Pentanoic acid, 4-oxo3.0.5.24.54.3 ± 1.1
Nonanoic acid1.02.52.31.9 ±0.8
Decanoic acid1.01.70.41.0 ± 0.6
Dodecanoic acid0.71.50.50.9 ± 0.5
Tetradecanoic acid7.29.95.17.4 ± 2.4
Hexadecanoic acid28.021.525.324.9 ± 3.3
9-Octadecenoic acid3.13.26.54.2 ± 1.9
Octadecanoic acid3.32.94.03.4 ± 0.5
Eicosanoic acid1.40.81.51.2 ± 0.4
Diacids
Octanedioc acid1.21.40.81.1 ± 0.3
Phenolics
Benzoic acid3.22.32.32.6 ± 0.5
Benzeneacetic acid1.40.90.81.0 ± 0.3
Syringol4.64.61.33.5 ± 1.9
Vanillin2.62.31.12.0 ± 0.8
Vanillin acid1.91.52.11.9 ± 0.3
Methyl p-coumarate 9.54.70.85.0 ± 4.4
Steroids
Stigmasterol1.30.51.31.1 ± 0.4
β-sitosterol6.82.95.14.9 ± 1.9
Sugars
Erythro-penitol, 2-deoxy-1,3,4,5 tetrakis-o-trimethylsilyl1.10.61.31.0 ± 0.4
meso-erythritol0.01.84.92.2 ± 2.5
Ribitol0.00.61.60.7 ± 0.8
Total identified compounds90.176.080.982.4 ± 7.2
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Pereira, H.; Simões, R.; Miranda, I. Cuticular Waxes and Cutin in Terminalia catappa Leaves from the Equatorial São Tomé and Príncipe Islands. Molecules 2023, 28, 6365. https://doi.org/10.3390/molecules28176365

AMA Style

Pereira H, Simões R, Miranda I. Cuticular Waxes and Cutin in Terminalia catappa Leaves from the Equatorial São Tomé and Príncipe Islands. Molecules. 2023; 28(17):6365. https://doi.org/10.3390/molecules28176365

Chicago/Turabian Style

Pereira, Helena, Rita Simões, and Isabel Miranda. 2023. "Cuticular Waxes and Cutin in Terminalia catappa Leaves from the Equatorial São Tomé and Príncipe Islands" Molecules 28, no. 17: 6365. https://doi.org/10.3390/molecules28176365

Article Metrics

Back to TopTop