Comparative Analyses of Cuticular Waxes on Various Organs of

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Comparative analyses of cuticular waxes on various organs of potato (Solanum tuberosum L.) Yanjun Guo, and Reinhard Jetter J. Agric. Food Chem., Just Accepted Manuscript • Publication Date (Web): 03 May 2017 Downloaded from http://pubs.acs.org on May 5, 2017

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Journal of Agricultural and Food Chemistry

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Comparative analyses of cuticular waxes on various organs of potato (Solanum

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tuberosum L.)

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Yanjun Guo1,2, Reinhard Jetter2,3

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1 College of Agronomy and Biotechnology, Southwest University, Chongqing,

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400716, China;

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2 Department of Botany, University of British Columbia, 6270 University Boulevard,

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Vancouver, BC V6T 1Z4, Canada;

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3 Department of Chemistry, University of British Columbia, 2036 Main Mall,

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Vancouver, BC V6T 1Z1, Canada;

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Corresponding author email address: [email protected]

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1

ACS Paragon Plus Environment


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ABSTRACT

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Complex mixtures of cuticular waxes coat plant surfaces to seal them against

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environmental stresses, with compositions greatly varying between species and

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possibly organs. This paper reports comprehensive analyses of the waxes on both

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above- and below-ground organs of potato, where total wax coverages varied between

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petals (2.6 µg/cm2), leaves, stems and tubers (1.8 - 1.9 µg/cm2), and rhizomes (1.1

22

µg/cm2). The wax mixtures on above-ground organs were dominated by alkanes,

23

occurring

24

2-methylalkanes and C26-C34 3-methylalkanes. In contrast, below-ground organs had

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waxes rich in monoacylglycerols (C22-C28 acyls) and C18-C30 alkyl ferulates, together

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with fatty acids (rhizomes) or primary alcohols (tubers). The organ-specific wax

27

coverages, compound class distribution and chain length profiles suggest highly

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regulated activities of wax biosynthesis enzymes, likely related to organ-specific

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ecophysiological functions.

in

homologous

series

of

isomeric

C25-C35

n-alkanes,

C25-C35

30 31

KEYWORDS: Biosynthesis; Chain length distribution; Cuticular wax; Solanum

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tuberosum (Potato); Wax esters.

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2


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INTRODUCTION

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Most plant organs are lined by hydrophobic surface structures to protect them

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from biotic and abiotic stresses, and it is well established that the primary function of

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these coatings is to seal tissues against excessive water loss.1, 2 However, they also

38

play important roles in mediating, e.g., herbivore and pathogen interactions, gas

39

exchange, photosynthetic and UV light reflection, organ development, and uptake of

40

xenobiotic chemicals.3 The various biological roles must be balanced for specific

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situations, and accordingly their functional performances have been found to vary

42

widely, as for example the cuticles lining leaf and fruit surfaces of diverse species

43

have water vapor permeabilities differing by up to two orders of magnitude.4 Such

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drastic differences in functional performance are thought to reflect differences in

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chemical composition of the surface coatings between respective plant species or

46

organs.

47

The cuticles lining above-ground plant surfaces are composite structures based

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on cutin, a polymer consisting mainly of ω- and mid-chain hydroxy or epoxy C16 and

49

C18 fatty acids and glycerol. 5 Within the cutin matrix, intracuticular wax is embedded,

50

and on the cutin surface further wax is deposited as an epicuticular layer.6 Due to their

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solubility in organic solvents, the cuticular waxes may be extracted directly from

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intact organs.7 Cuticular waxes are complex mixtures of very-long-chain fatty acids

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(VLCFAs) and their derivatives, including aldehydes, ketones, primary and secondary

54

alcohols, diols, ketols, diketones, alkyl esters and alkanes.8 Secondary metabolites,

55

such as triterpenoids, phenylpropanoids and flavonoids, occur in the wax mixtures of 3



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some plant species.8 Below-ground plant organs have internal as well as external

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transport barriers consisting of waxes embedded in suberin, a polyester similar to

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cutin but (depending on plant species and organs) also incorporating aromatic acids,

59

unsaturated fatty acids, or VLCFA components.9

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The biosynthesis of VLCFA wax constituents utilizes C16 fatty acyl precursors

61

produced in epidermal plastids, which are exported to the endoplasmic reticulum (ER)

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and activated to acyl-CoAs.10 Acyl elongation to VLCFAs is catalyzed by fatty acid

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elongase (FAE) complexes comprising a ketoacyl-CoA synthase (KCS), a

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ketoacyl-CoA reductase (KCR), a hydroxyacyl-CoA dehydratase (HCD) and an

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enoyl-CoA reductase (ECR). Other enzymes, including CER2-LIKEs and the

66

long-chain acyl-CoA synthetase CER8, are known to be involved in wax elongation in

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Arabidopsis thaliana,11 however their exact roles in the process are not understood.

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The acyl-CoAs resulting from FAE elongation are further modified along several

69

parallel pathways involving (1) head group reduction to alcohols and their

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esterification with fatty acids, (2) head group reduction to aldehydes, decarbonylation

71

to alkanes, and oxidation to secondary alcohols and ketones,10 and likely (3)

72

hydrolysis of the CoA thioesters to free fatty acids.

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The quantitative compositions of both cutin- and suberin-associated wax

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mixtures vary drastically between plant species and, in the few cases studied to date,

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also between organs and developmental stages.8,9,12 For example, the petal wax of

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Cosmos bipinnatus contains high concentrations of C22 and C24 fatty acids and

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primary alcohols, much shorter than those in leaf and stem waxes.13 The flag leaf 4


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blade wax of Triticum aestivum is dominated by primary alcohols, while peduncle

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wax comprises primarily β-diketones, suggesting differential regulation of the acyl

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reduction and β-diketone biosynthetic pathways in both organs.14 Below-ground

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organs contain aliphatic waxes typically differing from above-ground wax mixtures.15

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For example, Arabidopsis thaliana root waxes comprise primary alcohols, alkyl

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hydroxycinnamates, sterols and monoacylglycerols,16 while respective leaf wax is

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dominated by alkanes and primary alcohols, and stem wax by alkanes, secondary

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alcohols and ketones.17

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The waxes of relatively few crop species have been investigated systematically

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to date, notable exceptions including the leaf wax mixtures of various Brassica

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cultivars,18 the leaf, stem and spike waxes of diverse cereals,19-21 and the waxes on

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Rosaceae fruits such as apple, pear and cherry.22-24 In contrast, the waxes covering

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various parts of potato (Solanum tuberosum), one of the most important staple food

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crops on all continents, have been analyzed only sporadically. The few potato wax

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studies to date employed widely differing methods, and they focused on different

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cultivars and organs, thus impeding systematic comparisons and follow-up studies

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into the formation and biological functions of the waxes. Accordingly, various potato

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cultivars had different total amounts of leaf cuticular wax,25-27 all dominated by

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varying amounts of C25 to C33 n-alkanes, 2- and 3-methylalkanes. Besides, primary

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and secondary alcohols, fatty acids, aldehydes, alkyl esters, methyl ketones, sterols,

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β-amyrin, benzoic acid esters, and fatty acid methyl, ethyl, isopropyl and phenethyl

99

esters were reported.27 In the wound periderm of potato tubers, ferulate accumulation 5



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was observed together with suberin formation.28 However, comprehensive analyses of

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the wax mixtures coating other potato organs are missing to date. It is, therefore,

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currently not possible to assess the properties and functions of different potato organs,

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to address the genetic and biochemical mechanisms determining wax composition, or

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to inform targeted breeding approaches to modify wax composition and enhance crop

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performance.

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While early biochemical and genetic studies focused on the formation of select

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wax compounds in various crops, most of the molecular genetic investigations into the

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formation of waxes focused on the model species Arabidopsis thaliana. Only recently,

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first reports on the molecular genetics of wax biosynthesis in wheat were published,29

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partially confirming the previous findings for Arabidopsis thaliana but also revealing

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characteristic differences between the species. Based on these findings, molecular

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genetic investigations into the wax biosynthesis machinery of various other crops will

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be of interest, with great potential to understand the underlying regulatory

114

mechanisms and to assist breeding of stress-tolerant cultivars. To lay the foundation

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for such genetic studies, the current study aimed to provide comprehensive qualitative

116

and quantitative analyses of the cuticular waxes on all major organs of potato. To this

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end, waxes were sampled from mature leaves, stems, petals, rhizomes and tubers of

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the potato S. tuberosum cv. German Butterball, all major compounds were identified

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by GC-MS, and the total wax coverages on the potato organs as well as compound

120

class compositions, chain length profiles and isomer distributions were determined

121

using GC-FID. 6


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MATERIALS AND METHODS

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Plant materials and wax sampling

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Twelve plants of Solanum tuberosum cv. German Butterball were grown in a

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growth chamber, one per pot (30 cm diameter, 35 cm height) with 4 kg sterilized soil

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(Sunshine Mix4, SunPro), at 20°C /15°C in a long-day light cycle (14 h-day/10

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h-night) with 110 µE m-2 s-1 of photosynthetically active radiation. The plants were

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watered once a week during the first month and then twice a week during the second

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and third month. Leaves, stems and flowers were collected in the ninth week after

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sprouting. One fully extended leaf was cut from a node near the middle of each main

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stem, and one section (4-5 cm long) was cut from the adjacent main stem internode of

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each plant using tweezers and scalpel. Petals were manually separated from other

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flower parts using tweezers. Below-ground parts were harvested in the 12th week and

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rinsed under tap water, and the tubers (diameter > 3 cm; five tubers per individual)

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and rhizomes (4-5 cm long, three segments per individual) were separated using

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tweezers. Materials from three individuals were pooled into one sample, to yield four

137

independent biological replicates. All harvested plant parts were used immediately for

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wax extraction.

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Before wax extraction, photos of leaves, stems, petals and rhizomes were taken

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and subjected to pixel counting using the ImageJ software utility30 to determine

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surface areas. The volumes of the tubers were determined first by measuring the water

142

volume they displaced,31 and then their surface areas were calculated as Area = K

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(V)2/3, where V is the volume of the tuber (cm3) and K is a dimensionless constant set 7



144

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to 1.38 according to Houston.32

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Surface wax mixtures were extracted twice for 30 s with CHCl3, with volumes

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sufficient to cover the plant materials. The two extracts from each sample were then

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combined and filtered through glass wool, and the solvent was evaporated under N2.

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n-Tetracosane was added to the fresh plant material before extraction, 10 µg for

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leaves, stems and petal, 5 µg for rhizomes and tubers.

150 151

Wax sample preparation and GC analysis

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Wax samples were prepared for GC analysis by dissolving them in pyridine (20

153

µL, Aldrich), then adding bis-N,O-trimethylsilyltrifluoroacetamide (BSTFA, 20 µL,

154

Aldrich). Mixtures were incubated at 70ºC for 45 min, then excess reagents were

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evaporated under N2, and finally CHCl3 (200 µL) was added.

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For compound identification, GC samples were analyzed using a 6890N Network

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GC (Agilent) equipped with a HP-1 capillary column (Agilent, length 30 m, inner

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diameter 320 µm, 1 µm film thickness). Each sample was injected on-column into a

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flow of He (1.4 mL/min). The GC oven was held at 50ºC for 2 min, heated at

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40ºC/min to 200ºC, held at 200ºC for 2 min, heated at 3ºC/min to 320ºC, and held at

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320ºC for 30 min. Compounds were detected with an Agilent 5973N Mass

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Spectrometric Detector (EI 70 eV; m/z 50-800, 1 scan s-1). Compounds were

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identified by comparing their mass spectra with published data and authentic

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standards.

165

Alkyl ferulates were identified based on characteristic fragment combinations 8


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m/z 73, 179, 192, 209, 219, 236, 249 and 266, together with respective molecular ions

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m/z 518, 546, 574, 602, 630, 658 and 686.33 Isomeric alkanes were baseline-separated

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under the current GC conditions, with 2-methylalkanes (iso-alkanes) eluting first,

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3-methylalkanes (anteiso-alkanes) second, and unbranched alkanes (n-alkanes) third.

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Isomers were identified based on characteristic abundance ratios of MS fragments

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[M-15]+, [M-29]+, [M-43]+ and [M-57]+. SN-1 Monoacylglycerols (MAGs) were

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identified based on common characteristic fragments m/z 73, 103, 129

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[M-RCOOH-OTMSi]+, and 147, in separate GC peaks combined with fingerprint

174

ratios of fragments m/z 203, 205 [M-RCOOH]+ (but little and m/z 218) and a base

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peak [M-CH2OTMSi]+.16 Triacylglycerols (TAGs) were identified based on

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characteristic fragment combinations, with m/z 211, 227 and 383 showing the

177

presence of C14 acyl(s), and m/z 99, 115 and 495 the presence of C6 acyl(s).34 All

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fingerprint fragments of TAG isomers with C6 acyl groups esterified in the SN-2

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position of glycerol were found in substantial quantities, whereas the distinctive

180

fragment m/z 481 of the isomer with C6 acyl on SN-1 could not be detected. Thus, the

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TAGs were unambiguously identified as SN-2 C6 acyl isomers.

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Compounds were quantified by GC coupled to a flame ionization detector set at

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250ºC, burning H2 (30 mL/min) in air (200 mL/min), with the flame shaped by N2 (20

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mL/min). GC conditions were as for compound identification, but with H2 carrier gas

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(2.0 mL/min). Wax compound peak areas were compared against the internal standard

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peak area for quantification. The relative response factors relative to the internal

187

standard were taken as 1.00 for all cuticular wax constituents regardless of chain 9



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length and compound class, in agreement with past reports using the same GC

189

conditions. 7

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Statistical Analysis

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Data are presented as the means ± standard error of four independent samples.

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One-way ANOVA was used to compare the difference of total wax coverage among

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organs (SPSS 17.0, USA). The differences between means were evaluated using

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Tukey HSD tests. Statistical significance was considered at P < 0.05.

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RESULTS AND DISCUSSION

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Cuticular waxes were extracted from the various organs of S. tuberosum,

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identified by GC-MS and quantified by GC-FID. To comprehensively describe the

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wax mixtures, we determined the wax coverage and compound class composition,

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chain length distributions of fatty acids, alcohols and alkanes, and ester compositions.

201 202

Wax coverage and class composition

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All the potato organs were covered by similar wax amounts, with coverages

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varying slightly between petals (2.56 µg/cm2), leaves, stems and tubers (1.78 to 1.93

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µg/cm2), and rhizomes (1.14 µg/cm2) (Fig.1). The wax amounts found here were

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lower than in previous studies focusing on some of the potato organs, where for

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instance 5-7 µg/cm2 had been reported for potato leaves,26,27 possibly due to

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differences in cultivars or growing conditions.7,35 The coverage found here for tubers

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extracted immediately after harvest was substantially lower than the coverages of

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10-50 µg/cm2 previously reported for isolated tuber periderm, which increased with

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prolonged storage time within four weeks after harvest for both native periderm and

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wound periderm.33 It seems plausible that the potato tubers, functioning as long-term

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storage and reproductive organs, had higher wax coverages than the elongated

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internodes along the diageotropic shoots of the rhizomes.36,37

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The wax mixtures on the different potato organs all contained fatty acids,

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primary alcohols and n-alkanes (Fig. 2). Leaf wax comprised n-alkanes (52%),

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2-methylalkanes (21%), primary alcohols (12%), fatty acids (9%) and secondary 11



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alcohols (0.3%). Stem wax consisted of n-alkanes (47%) and 2-methylalkanes (9%),

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together with fatty acids, primary alcohols and TAGs (each ca. 14%). In the petal wax

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mixture, relatively large concentrations of n-alkanes (35%), 2-methylalkanes (43%)

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and 3-methylalkanes (19%) accumulated, accompanied by trace amounts of β-sitsterol.

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In contrast, the rhizome wax was dominated by fatty acids (54%), co-occurring with

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n-alkanes (5%), alkyl ferulates (13%), MAGs (9%), primary alcohols (5%), alkyl

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esters (2%) and β-sitsterol (1%). Tuber wax contained high amounts of alkyl ferulates

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(40%) and primary alcohols (36%,), accompanied by fatty acids (8%), MAGs (8%),

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n-alkanes (2%), β-amyrin (1%) and β-sitsterol (1%). Relatively small portions of the

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wax mixtures on all organs remained unidentified, ranging from 1% for rhizome wax

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to 4% for tuber wax.

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All the major compound classes identified here in potato leaf wax matched those

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detected in other studies.26,27 However, several other compounds that had been

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reported as relatively minor wax constituents could not be positively identified in the

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direct GC analyses of whole wax mixtures here. Most notably, neither methylketones

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nor aldehydes could be detected, even with the most sensitive detection restricted to

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single, characteristic MS fragments (m/z 58 for methyl ketones and m/z 82 or m/z 96

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for aldehydes). However, both compound classes had been identified before only after

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pre-separation of the potato leaf wax mixtures, at trace levels that varied greatly

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between cultivars.27

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The potato petal wax mixture, with relatively high concentration of alkanes, was

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similar to the petal waxes of Vicia faba and Antirrhinum majus.38,39 This is in contrast 12


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to other species, where for example Cosmos bipinnatus petal wax consisted mainly of

241

alcohols,13 and Petunia hybrid petal wax of VLCFA esters.40 Potato petal wax was

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further characterized by relatively high abundance of 2-methylalkanes and

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3-methylalkanes, reminiscent of previous reports on the occurrence of branched

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alkanes in petal waxes of Papaver rhoeas and Antirrhinum majus.38,41 It will be

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interesting to determine relative quantities of hydrocarbons, and of branched alkanes

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in particular, in the petal waxes of a broader range of plant species, and to correlate

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their occurrence with structural features such as crystallinity, surface morphology and

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distribution within the intra- and epicuticular wax layers in respective wax mixtures.

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Ultimately, petal wax variation may thus be explored to further our understanding of

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wax physiological and ecological performances,42,43 also in comparison between

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insect-pollinated and self-pollinated plant species (such as potato).

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Interestingly, relatively high amounts of alkyl ferulates were found in the potato

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rhizome and tuber waxes, and thus likely associated with suberized structures.44

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Bernards and Lewis28 reported that alkyl ferulate esters were restricted to the wound

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periderm, and their accumulation was shown to be correlated with the process of

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suberization in wound healing. Both alkyl ferulates and free primary alcohols were

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predominant in tuber wax, suggesting that high abundance of free alcohols

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contributed to the accumulation of alkyl ferulates, in accordance with previous studies

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showing that alkyl ferulates are formed by a potato fatty ω-hydroxyacid / fatty alcohol

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hydroxycinnamoyl transferase.45

13



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Overall, fatty acids and alcohols dominated the waxes on below-ground organs,

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whereas alkanes accumulated largely in above-ground organs (leaf, stem and petal).

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Our results thus suggest differential regulation of wax biosynthesis in different potato

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organs, leading to strongly enhanced flux through the alkane-forming pathway in the

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aerial organs and the acid/alcohol-forming pathways in tubers and rhizomes.

266 267

Chain length distributions of potato wax fatty acids, alcohols and alkanes

268

Most of the fatty acid-derived compound classes in the potato wax mixtures were

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present as homologous series, with substantial variance in chain length distributions

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between organs. Leaf wax contained a fairly broad range of unbranched fatty acids

271

(C20 to C30), similar to stem (C20 to C26), rhizome (C20 to C32) and tuber waxes (C22 to

272

C30). However, the potato organs had distinct quantitative profiles of fatty acids, with

273

the C26 homolog dominating on leaves, C24 on stems, and C28 on rhizomes and tubers

274

(Fig. 3). Even-numbered fatty acid homologs dominated on all organs, with minor

275

amounts of odd-numbered fatty acids observed mainly in rhizome wax.

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Only even-numbered, unbranched primary alcohols were detected, with chain

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lengths ranging from C22 to C34 in leaf wax, from C22 to C30 in stem wax, and from

278

C22 to C30 in rhizome and tuber waxes. The alcohol fractions of both leaf and stem

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waxes were dominated by the C26 and C28 homologs, while alcohol profiles peaked at

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C28 in tuber wax, and around C24 and C26 in rhizome wax. Only trace amounts of C30

281

fatty acid and C30 primary alcohol were detected in petal wax.

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n-Alkanes were observed in the wax mixtures extracted from all potato organs, 14


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with similar chain length ranges on leaf (C25 to C35), stem (C27 to C35), petal (C25 to

284

C33), rhizome (C25 to C31) and tuber (C25 to C29). Only odd-numbered homologs of

285

unbranched alkanes were detected, with profiles dominated by the C31 homologs on

286

leaves, stems and petals, and by C27 in rhizome and tuber waxes (Fig. 4). Leaf wax

287

contained 2-methylalkanes with odd carbon numbers ranging from C29 to C35 and

288

dominated by C33, from C29 to C34 (dominated by C31) in stem wax, and from C25 to

289

C35 (dominated by C29 and C31) in petal wax. 3-Methylalkanes with even carbon

290

numbers between C26 and C34 were identified in petal wax, with the C28 and C30

291

homologs dominating.

292

Among the three series of isomeric alkanes in potato waxes, both

293

2-methylalkanes and n-alkanes thus showed odd-over-even carbon number ratios,

294

whereas 3-methylalkanes exhibited the opposite even-over-odd preference. Similar

295

chain length distribution patterns of respective iso-branched, anteiso-branched and

296

straight-chain alkanes had also been observed, for example, in leaf waxes of

297

Nicotiana tabacum,46 and fruit waxes of Capsicum annuum and Solanum

298

melongena,47 as well as S. lycopersicum.48 Based on these parities and biochemical

299

evidence, it is generally accepted that the 2-methylalkanes and 3-methylalkanes

300

originate from iso-butyryl-CoA (C4) and 2-methylbutyryl-CoA (C5) precursors,

301

derived from valine and isoleucine, respectively. Elongation of these precursors in

302

plastidial FAS and ER-localized FAE complexes leads to even-numbered VLC

303

iso-acyls and odd-numbered VLC anteiso-acyls, analogous to the elongation of

304

acetyl-CoA (C2) starters to even-numbered straight-chain acyls. Loss of a C1 unit with 15



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305

the carboxyl function then yields 2-methylalkanes and n-alkanes with odd total carbon

306

numbers and 3-methylalkanes with even numbers. Our results on homolog

307

distributions of branched and unbranched potato wax alkanes are therefore in

308

accordance with the biosynthesis pathways established for other species.

309

The two below-ground potato organs had wax n-alkane chain length distributions

310

dominated by C27, coinciding with fatty acid profiles peaking at C28. Taken together,

311

these findings suggest that the n-alkane distributions reflected the abundance of their

312

fatty acyl precursors rather than substrate preference of enzymes along the

313

alkane-forming pathway (CER3 and CER1). In contrast, all three above-ground

314

organs had n-alkane fractions dominated by the C31 homolog while the (leaf and stem)

315

fatty acid and alcohol series peaked between C24 and C28, suggesting that the

316

alkane-forming enzymes here exerted substrate specificity for C32 and against C24 /

317

C26 acyl-CoAs. From this, it may be concluded that different CER3 and/or CER1

318

homologs may be operating in the above- and below-ground organs. Alternatively,

319

alkane formation may involve identical CER3 and CER1 homologs in all potato

320

organs, where they accept C28 – C32 acyl-CoAs in varying ratios depending on local

321

pools, but not C24 or C26 acyl-CoAs.

322 323

Chain length and isomer distributions of potato wax esters

324

Four different classes of esters were identified in the wax mixtures extracted

325

from various potato organs, in diverse combinations of aliphatic and aromatic acid

326

and alcohol moieties. All ester classes occurred in homologous series, in the fatty acid 16


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327

alkyl esters due to variation in both moieties, in the monoacylglycerides (MAGs) and

328

triacylglycerides (TAGs) due to variation of the acyl moiety, and in the alkyl ferulates

329

due to variation of the alkyl chain.

330

Potato rhizome wax contained fatty acid alkyl esters with total carbon numbers

331

of C42, C44 and C46 (Fig. 5). After initial GC-MS identification and GC-FID

332

quantification of these esters, a separate GC-MS experiment was carried out where

333

detection of diagnostic acylium fragments enabled the quantification of constituent

334

fatty acid moieties within each of these ester homologs. Only even-numbered fatty

335

acid moieties were observed in all three alkyl esters, with chain lengths ranging from

336

C12 to C26 in the C42 ester, from C16 to C30 in the C44 ester, and from C18 to C26 in the

337

C46 ester. The predominant acyl moiety varied, from C20 in the C42 ester to C22 in the

338

C44 ester and C24 in the C46 ester (Fig. 6). Accordingly, the alkyl esters contained only

339

even-numbered alcohol homologs, with the C22 alcohol dominating in all three ester

340

homologs.

341

Alkyl esters have been found in the cuticular waxes of many plant species, such

342

as Arabidopsis thaliana,49 Cyathea dealbata,50 Cereus peruvianus,51 Quercus robur,52

343

and Pinus radiata.53 In Arabidopsis thaliana, the wax esters are known to be formed

344

through esterification of primary alcohols and fatty acyl-CoAs, catalyzed by the

345

bifunctional wax ester synthase/diacylglycerol acyltransferase WSD1.54 Therefore, it

346

may be surmised that a WSD1 homolog may also be responsible for wax alkyl ester

347

biosynthesis in potato, where it is likely expressed preferentially in the rhizome.

17



Page 18 of 35

348

A series of TAGs was detected in potato stem wax, with total carbon numbers

349

ranging from C35 to C41 and the C37 homolog predominating (Fig. 5). All the TAGs

350

had C6 acyl units on the glycerol SN-2 position, and varying medium- to long-chain

351

acyls on SN-1/3. The C35 TAG thus comprised a combination of C12 and C14 acyls, the

352

C37 homolog two C14 acyls, the C39 homolog a combination of C14 and C16 acyls, and

353

the C41 homolog two C16 acyls. It is interesting to note that similar TAG compositions

354

had been reported for waxes of a few plant species before, for example in the leaf wax

355

of the distantly related Asteraceae species Cirsium arvense,34 but not for potato.

356

In potato rhizome and tuber waxes, a series of SN-2 MAGs with even fatty acids

357

ranging from C22 to C28 was detected, peaking at the C24 acyl homolog on both organs.

358

They were accompanied by lesser amounts of corresponding SN-1 isomers, with

359

nearly identical chain length range and distribution. Similar MAGs had been detected

360

in the waxes isolated from roots of Arabidopsis thaliana,16 roots of Ipomoea batatas,

361

Zea mays and Oryza sativa,15 and tuber periderm of potato,44 however with

362

predominantly SN-1 isomers. Such different results may be due to differences in

363

sample preparation, where migration of acyl groups to the thermodynamically favored

364

SN-1 position may occur,55 or else reflect product specificities of the esterifying

365

enzymes.56 In Arabidopsis thaliana, glycerol-3-phosphate acyltransferases (GPATs)

366

are involved in the synthesis of SN-1 and SN-2 MAGs, some of them accepting VLC

367

substrates and thus competing for the same pool of acyl-CoA substrates as cuticular

368

wax elongation/modification enzymes.57 Consequently, ectopic expression of GPAT5

369

led to the formation of saturated MAGs (and free fatty acids) as novel components of 18


Page 19 of 35


370

cuticular waxes.16 In this context it is of interest that the potato wax MAGs had acyl

371

chain length profiles largely diverging from those of the accompanying free fatty

372

acids (MAGs dominated by C24 acyl, free acids dominated by C28), suggesting that the

373

esterifying enzymes may show acyl chain length preference in this species.

374

Finally, potato rhizome and tuber waxes contained ferulic acid esters with both

375

even- and odd-numbered alcohol homologs ranging from C18 to C30. Even alcohols

376

dominated, peaking at C28 on both rhizomes and tubers (with 31% and 76%,

377

respectively). Similar alkyl ferulate compositions had been reported for root waxes

378

from Arabidopsis thaliana,16 Z. mays, O.sativa, Beta vulgaris and N. tabacum,15 as

379

well as for potato periderm.44 The potato alkyl ferulates associated with suberin are

380

known to be formed by BAHD family acyltransferases.58 In this context, the current

381

finding that the alkyl ferulate and free primary alcohols had similar chain length

382

profiles now suggests that (in potato) alkyl ferulate chain lengths are dictated by

383

substrate availability rather than transferase specificity.

384 385

In conclusion, this study provides comprehensive qualitative and quantitative

386

data on the suberin- and cutin-associated waxes on both above- and below-ground

387

organs of potato. Our analyses of potato leaf and tuber waxes confirmed previous

388

reports, while adding further information on compound class distributions, homolog

389

profiles and (ester) isomer compositions, in direct comparisons between both organs.

390

These findings can now be further compared with the compositions of potato stem,

391

petal and rhizome waxes, three organs that had not been analyzed before. All the 19



Page 20 of 35

392

potato organs showed similar cuticular wax coverages, but drastically different

393

compound class mixtures. The above-ground organs were dominated by alkanes

394

whereas the below-ground organs were dominated by esters (MAGs and acyl ferulates)

395

and fatty acids (rhizome) or primary alcohols (tuber). Branched alkanes were

396

observed in all above-ground organs, with 3-methylalkanes on petals only. The chain

397

length distributions of fatty acids and primary alcohols differed among organs, either

398

in free form or esterified TAGs or MAGs, ferulates or fatty acid alkyl esters.

399

Differences in the homolog and isomer distributions of various compound classes on

400

the different potato organs lead us to predict the existence of several wax biosynthesis

401

enzymes with distinct substrate specificities and differentially regulated activities in

402

various organs. The current data may thus serve as a foundation for future studies into

403

the molecular genetics and biochemistry of wax formation in potato.

20


Page 21 of 35


404

ABBREVIATIONS USED

405

BSTFA, bis-N,O-trimethylsilyltrifluoroacetamide; ER, endoplasmic reticulum; FID,

406

flame ionization detection; G3P, glycerol-3-phosphate; GPAT, glycerol-3-phosphate

407

acyltransferase; GC, gas chromatography; FHT, omega-hydroxy fatty acid/fatty

408

alcohol hydroxycinnamoyl transferase; KCS, 3-ketoacyl-CoA synthase; MAG,

409

monoacylglyceride; MAH, mid-chain alkane hydroxylase; MS, mass spectrometry;

410

PKS, polyketide synthase; TAG, triacylglyceride; VLCFA, very long chain fatty acid;

411

FAE, fatty acid elongase.

412 413

AUTHOR INFORMATION

414

Corresponding Author

415

Tel:+86 23 68251264. Email: [email protected]

416 417

ORCHID

418

Yanjun Guo

0000-0002-7252-3041

419 420

NOTES

421

The authors declare no competing financial interest.

422 423

ACKNOWLEDGMENTS

424

The authors thank Dr. Lucas Busta for his help with compound quantification and

425

identification. 21



Page 22 of 35

426 427

FUNDING SOURCES

428

Yanjun Guo conducted this study as a visiting scholar with financial support from

429

Chongqing Municipal Education Commission Fund. This research was funded by

430

National Natural Science Foundation of China (31670407), Chongqing Major Theme

431

Project (cstc2015shms-ztzx80004), and the Natural Science and Engineering Research

432

Council of Canada (Discovery grant #262461).

22


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433 434 435 436 437 438 439 440 441 442 443 444 445 446 447 448 449 450 451 452 453 454 455 456 457 458 459 460 461 462 463 464 465 466 467 468 469 470 471 472 473 474 475 476

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FIGURE CAPTIONS Figure 1. Wax coverages on the surfaces of various Solanum tuberosum organs. Data are given as averages of four biological replicates with standard errors. Different letters above the bars indicate significant differences between respective organs according to Tukey’s HSD tests (P

Comparative Analyses of Cuticular Waxes on Various Organs of

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