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Metabolic Changes during Storage of Brassica napus Seeds under Moist Conditions and Consequences for the Sensory Quality of the Resulting Virgin Oil Anja Bonte, Rabea Schweiger, Caroline Pons, Claudia Wagner, Ludger Brühl, Bertrand Matthäus, and Caroline Müller J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.7b04149 • Publication Date (Web): 05 Dec 2017 Downloaded from http://pubs.acs.org on December 11, 2017

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Journal of Agricultural and Food Chemistry

Metabolic Changes during Storage of Brassica napus Seeds under Moist Conditions and Consequences for the Sensory Quality of the Resulting Virgin Oil

Anja Bonte1,#, Rabea Schweiger2,#,*, Caroline Pons2, Claudia Wagner3, Ludger Brühl1, Bertrand Matthäus1, and Caroline Müller2

1

Department of Safety and Quality of Cereals, Max Rubner-Institut, Federal Research Institute of Nutrition and Food, Schützenberg 12, 32756 Detmold, Germany 2

Department of Chemical Ecology, Bielefeld University, Universitätsstr. 25, 33615 Bielefeld, Germany

3

Institute of Food Chemistry, University of Münster, Corrensstr. 45, 48149 Münster, Germany #

Both authors contributed equally

*Author for correspondence: Rabea Schweiger, [email protected], Phone: +49521-1065636

1 ACS Paragon Plus Environment


1 2

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ABSTRACT Virgin rapeseed (Brassica napus) oil is a valuable niche product, if delivered in high

3

quality. In this study, the effects of moist storage of B. napus seeds for one to four days

4

on the seed metabolome and the chemo-sensory properties of the produced oils were

5

determined. The concentrations of several primary metabolites including

6

monosaccharides and amino acids rapidly increased in the seeds, probably indicating

7

the breakdown of storage compounds to support seed germination. Seed

8

concentrations of indole glucosinolates increased with a slight time offset suggesting

9

that amino acids may be used to modify secondary metabolism. The volatile profiles of

10

the oils were pronouncedly influenced by moist seed storage, with the sensory quality of

11

the oils decreasing. This study provides a direct time-resolved link between seed

12

metabolism under moist conditions and the quality of the resulting oils, thereby

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emphasizing the crucial role of dry seed storage to ensure high oil quality.

14 15

KEYWORDS

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Brassica napus, seed, storage conditions, glucosinolates, oil, sensory quality


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INTRODUCTION Vegetable, edible oils are produced from several domestic crops with high seed oil

18 19

contents. These oils are important components of human nutrition, as they deliver

20

energy, improve the aroma of foods and are healthy due to high proportions of essential

21

unsaturated fatty acids in the triglycerides.1 Vegetable oils are classified according to

22

the plant species of origin and processing steps during oil production. Virgin vegetable

23

oils are produced by cold-pressing of seeds under mild conditions followed by

24

sedimentation or filtration of the oil, without further treatments of the seeds or the oil

25

(reviewed by Matthäus and Brühl2). By using such a gentle processing, nutritive and

26

healthy ingredients are retained and the oils have a more intense color, taste, and odor

27

compared to refined oils. However, studies are lacking that link metabolic processes in

28

the seeds with chemo-sensory properties of the corresponding oils, when seeds are

29

stored improperly. Virgin rapeseed (Brassica napus) oil is a valuable niche product on the European

30 31

market, as it contains high proportions of unsaturated fatty acids and further bioactive,

32

health-promoting compounds including vitamins, tocopherols, phenols, and flavonoids.3-

33

5

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fresh rapeseed, accompanied by a nutty after-taste.6 Brassica napus belongs to the

35

family Brassicaceae. Members of this taxon often contain erucic acid and glucosinolates

36

as prominent secondary plant compounds. Erucic acid is bitter and can cause heart

37

disease in animals and glucosinolates turn the seed cake (i. e., the protein-rich material

38

remaining after oil pressing) less useable as animal fodder. Therefore, today rapeseed

39

varieties with low erucic acid and glucosinolate concentrations (i. e., double-

It has a pleasant, characteristic mild aroma described with attributes like green, or



40

zero/double-low varieties) are used for the production of edible rapeseed oil. However,

41

glucosinolates are low in concentrations but not absent in these varieties. The sulfur-

42

containing glucosinolates and their volatile hydrolysis products are responsible for the

43

characteristic taste and odor of Brassicaceae crops, respectively.7-9 The volatile profile

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of high-quality virgin rapeseed oil is well characterized with its typical smell being

45

formed by a bouquet of diverse volatiles.10

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For acceptance of virgin rapeseed oil on the market and consumer satisfaction, a

47

reliable high sensory quality in terms of a characteristic odor and taste is of particular

48

importance.6 Indeed, virgin rapeseed oils with good sensory properties can be

49

distinguished from oils with poor sensory properties via sensory evaluation.10 Good oils

50

are characterized by the sensory attributes seed-like and nutty, sometimes

51

accompanied by astringent and strawy/woody, whereas poor oils are associated with

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off-flavors like musty/fusty, roasted/burnt, bitter, or rancid.6,10 Besides these sensory

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attributes, the oxidation stability and contents of several metabolites contribute to the oil

54

quality, which can be quite heterogeneous on the market.3

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As the oil quality cannot be improved after the pressing step, any adverse effects of

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seed management or processing leading to a reduced quality have to be avoided

57

making the production of high-quality virgin rapeseed oil quite challenging.2 Ideally, B.

58

napus seeds are stored at low moisture and low temperature to maintain weak

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metabolic rates and high seed viability but to prevent at the same time surface molds,

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seed germination, and adverse effects on oil/fat quality.11-14 High moisture probably is

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one of the most challenging problems for oil mills, especially if there is rain during

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harvest of the seed material. Under moist conditions, seeds may initiate germination4 ACS Paragon Plus Environment

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related metabolic processes and microbial communities may grow on seed surfaces.

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Both processes probably reduce the quality of the produced oils.

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A few studies describe the effects of seed storage under moist conditions and/or high

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temperatures on surface molds, seed germination, and oil/fat quality.11-13,15 However,

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little is known about the link between metabolic processes in the seeds and the chemo-

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sensory properties of the resulting oils. Moreover, impairments of oil quality caused by

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inappropriate seed storage may arise already within few days of moist conditions and

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probably depend on the duration of storage under these conditions. Thus, the aim of

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this study was to determine the effects of short-term (1-4 days) moist storage of B.

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napus seeds in a time-resolved manner on the seed metabolome and the chemo-

73

sensory properties of virgin oils produced from these seeds. We used a multi-platform

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metabolomics approach for comprehensively characterizing the seed and oil

75

metabolomes and linked these metabolic profiles to gain insight in various metabolic

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effects of moist seed storage relevant for oil quality.

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MATERIALS AND METHODS

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Chemicals and reagents. Chloroform (HPLC grade) was obtained from AppliChem

80

GmbH (Darmstadt, Germany). Methanol for GC-MS and UHPLC-FLD analyses (LC-MS

81

grade) was purchased from Fisher Scientific (Loughborough, UK). Methanol for HPLC-

82

DAD (HPLC grade) analyses was purchased from VWR International (Fontenay-sous-

83

Bois, France). Ribitol (99%), n-alkanes (C8-C40), octane, nonane, and undecane were

84

obtained from Sigma-Aldrich Chemie GmbH (Steinheim, Germany). L-Norvaline,

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sarcosine, ortho-phthaldialdehyde (OPA) reagent (10 mg mL-1 in 0.4 M borate buffer



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and 3-mercaptoproprionic acid), and 9-fluorenyl-methyl chloroformate (FMOC) reagent

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(2.5 mg mL-1 in acetonitrile) were purchased from Agilent Technologies (Santa Clara,

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CA, USA; Waldbronn, Germany). 2-Propenyl glucosinolate (sinigrin) was purchased

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from Phytoplan Diehm & Neuberger GmbH (Heidelberg, Germany). Sephadex (DEAE

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SephadexTM A-25) was obtained from GE Healthcare Bio-Sciences AB (Uppsala,

91

Sweden). Pentane, heptane, and tridecane were from Merck KGaA (Darmstadt,

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Germany). H2O was of Millipore grade. Authentic reference standards were from Sigma-

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Aldrich Chemie GmbH (Steinheim and Taufkirchen, Germany), Merck KGaA, Carl Roth

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GmbH + Co. KG (Karlsruhe, Germany), Macherey-Nagel GmbH & Co. KG (Düren,

95

Germany), Agilent Technologies, and H+R AG (Salzbergen, Germany).

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Storage experiment. Brassica napus seeds were obtained from a German oil mill.

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Seed material was a mixture of different double-low winter-sown rapeseed varieties

98

from conventional farming systems harvested in summer 2013. The storage experiment

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was conducted in August 2014. Before starting the experiment, the seeds had an initial

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moisture content of 7.6%. Subsamples of the seed material were taken as control

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samples (T0) for seed harvest and oil pressing (see below). The remaining seed

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material was placed on eight plates (47 x 30 x 2 cm; circa 300 g per plate) and daily

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humidified with tap water (ca. 500 mL per plate per day). The plates were stored at

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room temperature under daylight conditions. On four consecutive days (T1, T2, T3, T4),

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two of the plates (A, B) were taken for seed harvest and oil pressing. The time range of

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four days was used, because it is of practical relevance for oil mills. After four days

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germination is in an advanced state and such seeds will be refused by oil mills if

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germination is progressed too far.


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Within each plate, five subsamples of seeds (four in the edges and one in the middle

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of the plate) of about 2 mL were taken, immediately frozen in liquid nitrogen, and stored

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at - 80 °C until chemical analyses. This resulted in a sample number of n = 10 for each

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time point (2 plates x 5 subsamples). Circa 300 g of the remaining seeds of each plate

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were used to press oil resulting in two oils (A, B) per time point. After air-drying for 24 h

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(T1-T3) and 72 h (T4), respectively, the unpeeled seeds were pressed to oil using a

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screw extrusion press (Komet, IBG Monforts & Reiners GmbH & Co. KG,

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Mönchengladbach, Germany). Because oils cannot be pressed if the seed material is

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too moist or seeds are in an advanced state of germination, seed material was mixed

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with the same volume (1:1, v:v) of dry (T0 group) seeds. The oils were left for 24-72 h at

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4 °C for sedimentation, filtered with a glass fibre filter (13400-50-Q, Sartorius,

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Göttingen, Germany), filled in brown glass bottles, and stored at 4 °C until chemical

121

analysis.

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Seed metabolome. To comprehensively investigate the seed metabolome, a multi-

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analytical-platform metabolomics approach was chosen. Blanks (without plant material)

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were measured to discriminate between background- and sample-derived peaks.

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Authentic reference standards were used for metabolite identifications. Seeds were

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lyophilized for 48 h and the seed material of each sample was divided into subsamples

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for different analyses (one seed per subsample for GC-MS analyses; three seeds for

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UHPLC-FLD and HPLC-DAD analyses, respectively). The subsamples were ground,

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dry weights of the seed pellets determined (for normalization during data processing,

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see below), and the seed pellets slackened in a standardized manner (20 times each)

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with a dissecting needle to improve the extraction of metabolites.



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Metabolite profiling of carbohydrates, organic acids, and myo-inositol was performed

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according to Schweiger et al.16 using one seed per sample. Seed pellets were extracted

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in chloroform:methanol:H2O containing ribitol as internal standard. Aqueous phases

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were dried, methoximated, and silylated. Samples and n-alkanes (C8-C40) were

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analyzed via gas chromatography coupled to mass spectrometry (GC-MS; Focus GC,

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DSQII MS; Thermo Electron, Rodano, Italy) using a VF-5 ms column (30 m, 0.25 mm

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i.d.; Varian, Palo Alto, CA, USA) and electron impact (EI) positive ionization at 70 eV

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(full scan, 50-750 m/z). Analytes were identified by comparing Kováts retention indices17

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(RIs) and mass spectra with entries in the Golm metabolome database18 and authentic

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reference standards. For quantification, peak areas of analytes of the same metabolite

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were added together, related to those of ribitol and the seed pellet dry weights. Due to

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technical problems, sample sizes of the T1 and T3 group were reduced to n = 9.

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Amino acid profiling was done as described in Jakobs and Müller19 using three seeds

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per sample. Seed pellets were extracted with 80% methanol containing norvaline and

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sarcosine as internal standards and extracts filtered through 0.2 µm

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polytetrafluorethylene filters (Phenomenex, Torrance, CA, USA). Samples were

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analyzed via ultra-high performance liquid chromatography coupled to fluorescence

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detection (UHPLC-FLD; 1290 UHPLC, 1260 FLD, Agilent Technologies) after pre-

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column derivatization with OPA and FMOC. Amino acids were separated on a ZORBAX

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Eclipse Plus C18 column (250 mm x 4.6 mm i.d., 5 µm, Agilent Technologies) using the

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gradient described in Jakobs and Müller19 and detected via FLD. Amino acids were

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identified by comparing their retention times with those of authentic reference

154

standards. They were quantified as peak areas, which were related to those of the


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internal standards (norvaline for primary, sarcosine for secondary amino acids) and the

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seed pellet dry weights.

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Glucosinolate profiling was done as described in Abdalsamee and Müller20 using three

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seeds per sample. Seed pellets were extracted in 80% methanol, adding 2-propenyl

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glucosinolate (sinigrin) as internal standard. Glucosinolates were converted to

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desulfoglucosinolates on ion-exchange diethylaminoethyl Sephadex A-25 columns

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using sulfatase as described20 and analyzed using HPLC coupled to diode array

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detection (HPLC-DAD, 1200 Series, Agilent Technologies) equipped with a Supelcosil

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LC-18 column (250 mm x 4.6 mm, 5 µm, Supelco, PA, USA) using the gradient as

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described20. Glucosinolates were identified by comparing retention times and UV

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spectra with entries in an internal database, which had been confirmed by LC-MS.21,22

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For quantification, peak areas at 229 nm were related to the amount of the internal

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standard sinigrin, response factors for glucosinolates23, and seed pellet dry weights.

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Across all analytical platforms, only those metabolites, which were found in at least

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half of the samples of at least one treatment group, were kept for further data analyses.

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Total glucosinolate concentrations were compared between treatment groups using a

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Kruskal-Wallis test in R,24 as data were not normally distributed (tested with Shapiro-

172

Wilk test). After replacing zero values by small (10-13-10-12) random numbers as well as

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mean-centered unit-variance scaling (i.e., auto-scaling) of the data, principal component

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analyses (PCA) were conducted in R separately for data derived from the different

175

analytical platforms. For each metabolite, fold changes for the groups T1, T2, T3, and

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T4 were computed as mean metabolite pool sizes in these groups divided by mean pool

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sizes in the T0 group. Fold changes were log2-scaled for symmetry around zero. If there



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was a valid qualitative difference (i. e., the metabolite was found in at least half of the

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samples of one group and absent in the other group), the fold change was set to the

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minimum (decrease in pool size compared to T0) or maximum (increase in pool size)

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fold change observed in the whole dataset, respectively. If the metabolite was found in

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less than half of the samples of one group and was absent in the other group, this was

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interpreted as qualitative difference by chance and hence no fold changes were

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computed. Pool sizes of metabolites were regarded as considerably lower in the

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treatment (compared to the T0) group if fold changes were < 0.5 (log2 scale: < -1) and

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higher if fold changes were > 2 (log2 scale: > 1). Fold changes were plotted as heatmap

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stripes using Cluster 3.025 and Java TreeView 1.1.6r426 and mapped on a KEGG

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PATHWAY-derived27 metabolic map adjusted after Schweiger et al.16

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Volatile profile of the resulting oil. The volatile compounds of the oils were

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determined via dynamic headspace GC-MS modified according to Bonte et al.10 Each

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oil was measured five times over five consecutive days (starting 14-18 d after pressing

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the oils at T0) to test whether the chemical profile of the oils changes over time. In

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between, oils were stored at 4 °C. Analysis was carried out using the method DGF-C-VI

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20(15).28 Oils were mixed with alkanes (pentane, heptane, octane, nonane, undecane,

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tridecane) for RI calculation and heated to 80 °C in a PTA 3000 dynamic headspace

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system (IMT, Vohenstrauß, Germany). Purged volatiles were trapped, transferred to a

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GC (Trace 1300 Series, Thermo Scientific, Darmstadt, Germany), separated on a CPSil

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19 fused silica capillary column (14% cyanopropyl-phenyl / 86% dimethylpolysiloxane,

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60 m x 0.32 mm, 1 µm film thickness), and ions (positive EI mode, 35-300 m/z) detected

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in a ISQ single quadrupole mass spectrometer (Thermo Scientific). Metabolites were


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identified via comparison of mass spectra with the NIST Chemistry WebBook

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(http://webbook.nist.gov)29 and comparison of RIs and mass spectra with authentic

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reference standards. Quantification was done with the MeltDB platform.30 TAGs were

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detected automatically, if occurring in 80% of all chromatograms or in 80% of the

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samples of one group. In contrast to the metabolite profiling of seeds (see above),

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unidentified features were also quantified, as they may contribute to the sensory

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impression of the oils. A PCA was conducted and fold changes were computed as

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described above.

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Sensory quality of the oil. The sensory oil quality was assessed by a sensory panel

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of four trained persons according to the standard method of the German Society for Fat

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Science (DGF) C-II 1.28 The occurrences of the characteristic rapeseed oil attributes

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seed-like, nutty, strawy/woody, and astringent as well as the off-flavors roasted/burnt,

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bitter, rancid, musty/fusty, and others were determined using a scale from 0 (not

214

perceived) to 5 (intense). The testers evaluated the two replicate oils (A/B) per

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treatment group presented in blue-colored glasses with lids individually in a room that

216

was exclusively used for sensory evaluations at 20-22 °C. For each attribute, the

217

median intensity (across the four panellists) was calculated and then the means of the

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two medians of the replicate oils (A/B) of each treatment group were taken.

219 220 221 222

RESULTS After one day under moist conditions, some seed coats were already burst and seeds rapidly germinated within some days (Figure 1). At four days after moist storage,



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expanding cotyledons were clearly visible. The proportion of germinated seeds (across

224

plates A, B) steeply increased over time.

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In the seed extracts, 29 primary metabolites could be identified (Table 1). Compared to

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the control group T0, the primary metabolite profiles of seeds stored under moist

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conditions were shifted already after one day of moist storage, with the strongest

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metabolic shifts being obvious at three to four days after moist storage (Figures 2A-D).

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These metabolic shifts were mainly due to enhanced metabolite pool sizes of glucose,

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fructose, glyceric acid, dehydroascorbic acid, myo-inositol, and most amino acids

231

(Figures 2A-D, 3). In contrast, raffinose (and, to a lower extent, citric acid) showed

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decreased pool sizes after storage under moist conditions (Figures 2A-D, 3). As

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secondary metabolites, 13 glucosinolates were found in the seed extracts (Table 1).

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The total glucosinolate concentrations ranged from 2.4 to 19.4 µmol g-1 dry weight. By

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trend, glucosinolate concentrations increased with the moist seed storage time, but the

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differences were not significant (data not shown; Kruskal-Wallis test: Χ² = 8.361, df = 4,

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p = 0.079). The seed glucosinolate profiles were strongly affected by moist storage

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(Figures 2E-F). Compared to the changes in the pool sizes of several primary

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metabolites, the shift in glucosinolate composition occurred later. The glucosinolate

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profiles of (germinated) seeds of the T3 and T4 group were clearly separated from the

241

T0 and T1 group, whereas the profiles of the T2 group were in between. This metabolic

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shift, which started after two days of moist storage, was mainly attributable to an

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increase in concentrations of indole glucosinolates and, to a lesser extent, the benzyl

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glucosinolate gluconasturtiin, whereas aliphatic glucosinolates were unaffected or even

245

decreased (Figures 2E-F, 3).


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In the headspace of the virgin rapeseed oils, 60 metabolites of several substance

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classes could be identified (Table 2) and further 27 unidentified TAGs were detected

248

(Figure S1). The volatile profiles of the oils from seeds, which were stored for different

249

time spans under moist conditions, could be clearly distinguished (Figure 4A).

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Treatment effects were much stronger than metabolic shifts within the oils over time, as

251

assessed by repeatedly measuring oils over five consecutive days. Some metabolic

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features were slightly lower concentrated in the oils after moist seed storage for one to

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two days (T1/T2 versus T0 group; Table 2, Figure 4B, Figure S1) compared to the oils

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pressed at T0. However, the highest loadings and fold changes were found for

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compounds having higher concentrations in oils from seeds stored for four days under

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moist conditions (T4) compared to the other groups (Table 2, Figure 4B). Besides

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alcohols (3- and 2-methylbutan-1-ol), aldehydes (2-methylpropanal, 2-methylbutanal),

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and organosulfur compounds (carbonylsulfide, dimethyldisulfide, dimethyltrisulfide), all

259

isothiocyanates and some nitriles detected in this study were increased in the oils of the

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T4 group (Table 2, Figure 4B).

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The sensory description of the resulting native oils of the rapeseed samples changed

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with increasing seed storage time under moist conditions. The control oil pressed from

263

dry seeds (T0) was described with the characteristic attributes of a good virgin rapeseed

264

oil, i.e., seed-like and nutty6,10 (Figure 5). Some testers additionally assigned the

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attribute strawy/woody, which can occur without indicating something negative.6 No off-

266

flavors were assigned. On average, for those oils pressed from seeds stored under

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moist conditions for one to four days (T1-T4) the positive attributes seed-like and nutty

268

were assigned with lower intensities, whereas the intensities of the attributes



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strawy/woody and astringent were higher. Other (atypical) impressions occurred in

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those oils pressed from moist seeds only (Figure 5). The testers described these other

271

impressions as germinated, horseradish-like, spicy, and cress-like, respectively. The

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impression germinated in the oils appeared and increased from the second day of moist

273

seed storage on, being detected by none (T0, T1), two (T2, T3), and all four (T4) testers

274

consistently in both replicate (A/B) oils (data not shown).

275 276

DISCUSSION

277

For the acceptance of virgin rapeseed oil on the market and consumer satisfaction, a

278

reliable high oil quality is crucial. To understand how this quality is influenced by moist

279

seed storage, this study aimed in linking metabolic processes in the seeds with chemo-

280

sensory properties of the corresponding oils in a time-resolved manner.

281

In general, whereas some of the seed and oil metabolites may be of plant origin,

282

others could be produced by microorganisms associated with the seed material used for

283

oil pressing. Indeed, bacteria are known to produce various metabolites of diverse

284

substance classes.31 Because moisture induced seed germination (Figure 1) but

285

probably also induced growth and metabolic activities of the associated microbiome,

286

metabolic changes in the seeds and oils may be caused by both the plants and the

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microorganisms. However, it is difficult to ascribe the production of certain metabolites

288

to plant versus microorganism metabolism and for the overall sensory quality of the oil

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the source is not relevant.

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The primary metabolites detected in the seeds (Table 1) mainly matched those found

291

earlier in B. napus seeds.32 The rapid increase in concentrations of most seed primary


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metabolites under moist conditions (Figures 2, 3) is probably due to a breakdown of

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storage compounds. The dominant storage compounds in mature B. napus seeds are

294

proteins and oils.33 Thus, the increase in concentrations of most amino acids is likely

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caused by a breakdown of storage proteins. Moreover, triglycerides were probably

296

degraded to fatty acids, because increased seed free fatty acid concentrations have

297

been found in B. napus seeds stored under moist conditions.13,14,34 As starch is only

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transiently accumulated during seed development but almost absent in mature embryos

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and seeds of rapeseed,33,35 the increases of glucose and fructose may be due to a

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breakdown of oligosaccharides. Indeed, pool sizes of raffinose were decreased after

301

moist storage. The moisture-induced increases of dehydroascorbic acid may be related

302

to increasing ascorbic acid concentrations during rapeseed germination reported

303

earlier.36 Overall, the rapid seed metabolic reprogramming likely represents the

304

transition from seed dormancy to active metabolism facilitating germination and

305

seedling growth.

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Most of the glucosinolates found in the seeds (Table 1) already have been reported for

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B. napus seeds.37-39 Consistently across studies, the aliphatic glucosinolates progoitrin

308

and gluconapin were among the dominant glucosinolates in (ungerminated) seeds (data

309

not shown).37-39 Compared to the primary metabolites, the moisture-induced shift in the

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glucosinolate profiles occurred one to two days later (Figures 2, 3). As glucosinolates

311

are biosynthesized from glucose and amino acids, the time offset may represent the

312

time needed for sufficient accumulation of these precursors (Figures 2, 3), redirection of

313

metabolic fluxes, and glucosinolate biosynthesis. Remarkably, whereas the total

314

glucosinolate concentrations only slightly increased under moist conditions, the



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concentrations of indole glucosinolates pronouncedly increased. This is consistent with

316

earlier studies showing that concentrations of aliphatic glucosinolates decreased,

317

whereas those of indole glucosinolates increased in seeds of B. napus during

318

germination.37,38 Whether the total concentration of glucosinolates decreases (like

319

reported earlier),37,38 increases (current study), or remains constant during germination

320

may depend on the degree of resource limitation in the seeds, as resources derived

321

from glucosinolate catabolism may be used if needed.38 Dormant B. napus seeds

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probably are well defended due to their hard seed coat and higher glucosinolate

323

concentrations compared to leaves.39 Indeed, glucosinolates and their hydrolysis

324

products play a pivotal role in plant defense against generalist plant antagonists.8

325

However, after the germination-induced burst of the seed coat, the embryos and

326

emerging seedlings are prone to attacks by herbivores or pathogens. The shift to higher

327

proportions of indole glucosinolates during germination found in the current study for B.

328

napus and for Arabidopsis thaliana earlier23 may confer enhanced resistance to

329

generalist plant antagonists, as the degradation products of indole glucosinolates are

330

particularly biologically active,40 but this has to be tested further.

331

Moist storage of the seeds was also mirrored in the oil volatile profiles. Many of the

332

compounds detected in the oil headspaces (Table 2) have also been found in earlier

333

studies in virgin rapeseed oils.10,41,42 Several of the alcohols, aldehydes, and ketones

334

may be derived from amino acids via Strecker degradation and related pathways or

335

from unsaturated fatty acids via oxidation.43-45 Moreover, dimethylsulfide,

336

dimethyldisulfide, and dimethyltrisulfide could be produced via Strecker degradation

337

from methionine.46 Nitriles and isothiocyanates are typical degradation products of


Page 17 of 46


338

glucosinolates8 and were probably formed via non-enzymatic or myrosinase-mediated

339

hydrolysis of the seed glucosinolates during oil pressing. Many of the volatiles showed

340

increased concentrations when seeds were stored under moist conditions, especially

341

after four days (Table 2, Figure 4). However, total concentrations of volatiles did not

342

significantly differ between groups (data not shown). In contrast to this finding, other

343

studies reported higher total concentrations of volatile compounds in sensory poor

344

compared to good virgin rapeseed oils.10,15 The increased oil concentrations of 2-

345

methylpropanal after moist seed storage fit well to the finding that this compound is

346

generally higher in poor compared to good virgin rapeseed oils.10 The increase of this

347

aldehyde may be linked to the increased seed amino acid concentrations under moist

348

storage, as it may be produced from amino acids (see above). The increases in

349

dimethyldisulfide and dimethyltrisulfide probably are related to higher availabilities of

350

their precursor, methionine (Figure 3).46 However, dimethylsulfide was only slightly

351

influenced by moist seed storage in the current study (Table 2, Figure 4) and higher in

352

sensory good rapeseed oils compared to those with a musty/fusty off-flavor in an earlier

353

study.47 The increased concentrations of all isothiocyanates (and some nitriles) are

354

assumed to be related to metabolic shifts in the seed glucosinolate profiles (Figures 2,

355

3). Strikingly, 4-isothiocyanato-1-butene was higher in sensory good compared to

356

sensory poor virgin rapeseed oils in an earlier study,10 but increased in oil prepared

357

after moist seed storage in the current study (Table 2, Figure 4). Concentrations of seed

358

indole glucosinolates increased under moisture (Figures 2, 3), but the isothiocyanates

359

and nitriles produced from these glucosinolates are unstable and rapidly converted to



360

other products.40 Thus, the link between changes in the seed glucosinolates and oil

361

isothiocyanates is somehow vague and needs further investigation.

Page 18 of 46

362

The slightly lower concentrations of aldehydes (e.g., butanal, pentanal, heptanal,

363

octanal) in oils pressed from seeds stored under moist conditions for one day (T1)

364

compared to those pressed from dry (T0) seeds (Table 2, Figure 4) were unexpected.

365

These aldehydes are known as degradation products from autoxidation of unsaturated

366

fatty acids45 and related to certain off-flavors in virgin edible oils if occurring in higher

367

concentrations (Table 2). However, despite this metabolic shift the T1 oils kept their

368

seed-like sensory impression (Figure 5). Given the well-known aroma-activity of

369

aldehydes (Table 2) and the fact that clear atypical sensory impressions occurred later

370

in parallel to increasing isothiocyanate concentrations (Figures 4, 5), these results

371

indicate the dominant impact of glucosinolate degradation products on the flavor of

372

rapeseed oil.

373

The shifts in the oil volatile profiles led to distinct sensory impressions. Many of the

374

rapeseed oil volatiles were described as odor- and aroma-active with compound-

375

specific odor thresholds (Table 2).10,42,48-50 Shifts in the composition of these volatile

376

metabolites may explain the increase of atypical sensory impressions of the oils along

377

with moist seed storage (Figure 5). In fact, chemical and aroma profiles of rapeseeds

378

and the oils pressed from these are quite correlated.47 Although neither off-flavor

379

attributes related to improper seed storage (i. e., musty/fusty), seed pre-pressing

380

treatment (roasted/burnt), or oil storage (rancid)10 were assigned to any oil, other

381

atypical sensory impressions for oils pressed from moist seeds (germinated,

382

horseradish-like, spicy, cress-like) indicate that increased isothiocyanate concentrations


Page 19 of 46


383

may be responsible for these sensory impressions. Indeed, isothiocyanates confer

384

sulfury, garlic, and pungent sensory impressions.48 Specifically, the increased

385

concentration of 4-isothiocyanato-1-butene (odor threshold: 5 mg kg-1, unpublished

386

data) probably contributes to the sensory poor impression of the corresponding oils, as

387

high contents of this volatile are associated with strong off-flavors.43 Although

388

glucosinolate degradation products to a certain degree confer the characteristic flavor

389

and pungency to rapeseed oils, they can also cause off-flavors. In contrast to germ oil,

390

germination-related sensory impressions found in this study (germinated, horseradish-

391

like, spicy, cress-like) after moist storage of the seeds are not desired for virgin

392

rapeseed oil and therefore occur as atypical off-flavors. Moreover, other oil volatiles

393

may contribute to the sensory impressions, with or without being aroma-active

394

themselves.10

395

Apart from these sensory impressions, which strongly affect oil acceptance by the

396

consumers, the actual implications of changes in oil quality for human nutrition and

397

health have to be considered. Moist storage of B. napus seeds lowers the proportion of

398

essential (poly)unsaturated fatty acids11,12 as well as the contents of tocopherols34 in

399

rapeseed oil, thereby reducing its value in terms of nutrition and health. The implications

400

of increased concentrations of indole glucosinolates in the seeds and isothiocyanates in

401

the oils are less obvious. These compounds can be anti-nutritional, toxic, and

402

goitrogenic, but can also prevent cancer and be anti-inflammatory, thereby promoting

403

health.5,9,51 Further studies are needed to assess the values of different virgin rapeseed

404

oils in terms of nutrition and health.



405

Page 20 of 46

In conclusion, this study demonstrates that moisture during B. napus seed storage

406

induces rapid metabolic shifts in the seeds, which translate into shifts in the volatile

407

profiles of the corresponding oils leading to atypical and undesirable sensory

408

impressions. By linking metabolic effects of moist seed storage on the seeds and

409

corresponding oils, this study adds significant knowledge to the field of virgin rapeseed

410

oil quality control. Specifically, it contributes to the understanding of the role of seed

411

germination for oil quality. Our study cannot uncover effects of long-term moist storage

412

of seeds, but emphasizes that even short moist seed storage of up to four days

413

pronouncedly impairs oil quality. This is of high relevance, because seeds stored for

414

longer time under moist conditions are probably not used for oil pressing due to visible

415

advanced germination and mold. However, seeds stored moistly for only some days like

416

in this study may be used for oil production, especially if they are mixed with dry seeds.

417

As the described metabolic shifts are irreversible,2 it is of utmost importance to ensure

418

proper seed storage conditions to produce high-quality virgin rapeseed oil in terms of

419

sensory impressions and value for nutrition and health.

420 421

ABBREVIATIONS USED

422

FMOC, 9-fluorenyl-methyl chloroformate; GC-MS, gas chromatography coupled to mass

423

spectrometry; HPLC-DAD, high performance liquid chromatography coupled to diode

424

array detection; m/z, mass-to-charge ratio; OPA, ortho-phthaldialdehyde; PC, principal

425

component;

426

chromatography coupled to fluorescence detection; abbreviations of metabolites: see

427

Tables 1, 2.

RI,

retention

index;

UHPLC-FLD,

ultra-high

performance

liquid


Page 21 of 46


428

ACKNOWLEDGMENTS

429

We thank Karin Djendouci for practical help with the glucosinolate analyses and the

430

Bioinformatics Resource Facility (BRF), which is part of the Center for Biotechnology

431

(CeBiTec) at Bielefeld University, for the expert technical support with the MeltDB 2.0

432

platform.

433

ASSOCIATED CONTENT

434

Includes mass spectra of unidentified TAGs of the volatiles from the headspaces of

435

rapeseed oils (PDF).

436

AUTHOR INFORMATION

437

Corresponding Author

438

*Phone: +49521-1065636; Fax: +49521-1062963; Email: rabea.schweiger@uni-

439

bielefeld.de

440

ORCID

441

Rabea Schweiger: 0000-0001-8467-4966

442

Funding

443

This IGF Project of the FEI is supported via AiF within the programme for promoting the

444

Industrial Collective Research (IGF) of the German Ministry of Economics and

445

Energy(BMWi), based on a resolution of the German Parliament.

446

Notes

447

The authors declare no competing financial interest.

448

Author Contributions



Page 22 of 46

449

AB, RS, LB, BM, and CM designed the study. RS, CP, and CM carried out the seed

450

metabolome analyses. AB, CW, LB, and BM did the chemical and sensory analyses of

451

the oils. AB, RS, CP, and CW analyzed the data. AB, RS, LB, BM, and CM wrote the

452

publication.

453


Page 23 of 46


454

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455

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moisture conditions. Pol. J. Natur. Sc. 2013, 28, 485-499.

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Soild-Phase Microextraction Use for the Characterization of Volatile Compounds

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Schweiger, R.; Baier, M. C.; Persicke, M.; Müller, C. High specificity in plant leaf metabolic responses to arbuscular mycorrhiza. Nat. Commun. 2014, 5, 3886.


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Reference Database Number 69; National Institute of Standards and

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Technology: Gaithersburg MD.

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Kessler, N.; Neuweger, H.; Bonte, A.; Langenkämper, G.; Niehaus, K.;

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Nattkemper, T. W.; Goesmann, A. MeltDB 2.0 – advances of the metabolomics

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software system. Bioinformatics 2013, 29, 2452-2459.

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Schulz, S.; Dickschat, J. S. Bacterial volatiles: the smell of small organisms. Nat.

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Sinapine Esterase BnSCE3 in Oilseed Rape Seeds Triggers Global Changes in

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Seed Metabolism. Plant Physiol. 2011, 155, 1127-1145.

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Norton, G.; Harris, J. F. Compositional Changes in Developing Rape Seed

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in Tocochromanol Content in Seeds of Brassica napus L. During Adverse

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Conditions of Storage. J. Am. Oil Chem. Soc. 2011, 88, 1379-1385.

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da Silva, P. M. F. R.; Eastmond, P. J.; Hill, L. M.; Smith, A. M.; Rawsthorne, S.

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Starch metabolism in developing embryos of oilseed rape. Planta 1997, 203,

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rapeseed cultivars. Plant Foods Hum. Nutr. 1995, 47, 63-70.

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Glucosinolates in Germinating Seeds of White Mustard, Red Radish, White

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Radish, and Rapeseed. J. Agric. Food Chem. 2008, 56, 9087-9093.

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Clossais-Besnard, N.; Larher, F. Physiological Role of Glucosinolates in Brassica

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napus. Concentration and Distribution Pattern of Glucosinolates among Plant

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Organs during a Complete Life Cycle. J. Sci. Food Agric. 1991, 56, 25-38.

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Velasco, P.; Soengas, P.; Vilar, M.; Cartea, M. E; del Rio, M. Comparison of

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Glucosinolate Profiles in Leaf and Seed Tissues of Different Brassica napus

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Crops. J. Am. Soc. Hortic. Sci. 2008, 133, 551-558.

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Agerbirk, N.; de Vos, M.; Kim, J. H.; Jander, G. Indole glucosinolate breakdown and its biological effects. Phytochem. Rev. 2009, 8, 101-120.



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Hu, W.; Zhang, L.; Li, P.; Wang, X.; Zhang, Q.; Xu, B.; Sun, X.; Ma, F.; Ding, X.

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Characterization of volatile components in four vegetable oils by headspace two-

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dimensional comprehensive chromatography time-of-flight mass spectrometry.

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Talanta 2014, 129, 629-635.

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Matheis, K.; Granvogl, M. Characterisation of the key aroma compounds in

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commercial native cold-pressed rapeseed oil by means of the Sensomics

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approach. Eur. Food Res. Technol. 2016, 242, 1565-1575.

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Zhou, Q.; Yang, M.; Huang, F.; Zheng, C.; Deng, Q. Effect of Pretreatment with

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Dehulling and Microwaving on the Flavor Characteristics of Cold-Pressed

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Rapeseed Oil by GC-MS-PCA and Electronic Nose Discrimination. J. Food Sci.

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Yaylayan, V. A. Recent Advances in the Chemistry of Strecker Degradation and

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Amadori Rearrangement: Implications to Aroma and Color Formation. Food Sci.

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Choe, E.; Min, D. B. Mechanisms and Factors for Edible Oil Oxidation. Compr.

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Ballance, P. E. Production of volatile compounds related to the flavour of foods

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from the Strecker degradation of DL-methionine. J. Sci. Food Agric. 1961, 12,

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532-536.

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Matheis, K.; Granvogl, M. Characterization of Key Odorants Causing a

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Fusty/Musty Off-Flavor in Native Cold-Pressed Rapeseed Oil by Means of the

584

Sensomics Approach. J. Agric. Food Chem. 2016, 64, 8168-8178.


Page 29 of 46

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Engel, E.; Baty, C.; le Corre, D.; Souchon, I.; Martin, N. Flavor-Active

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Compounds Potentially Implicated in Cooked Cauliflower Acceptance. J. Agric.

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Food Chem. 2002, 50, 6459-6467.

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Pollner, G.; Schieberle, P. Characterization of the Key Odorants in Commercial

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Cold-Pressed Oils from Unpeeled and Peeled Rapeseeds by the Sensomics

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Approach. J. Agric. Food Chem. 2016, 64, 627-636.

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Related Compounds. J. Agric. Food Chem. 1999, 47, 4825-4836.

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Cartea, M. E.; Velasco, P. Glucosinolates in Brassica foods: bioavailability in food and significance for human health. Phytochem. Rev. 2008, 7, 213-229.

594 595

Sablé, S.; Cottenceau, G. Current Knowledge of Soft Cheeses Flavor and

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Díaz-Maroto, M. C.; Díaz-Maroto Hidalgo, I. J.; Sánchez-Palomo, E.; Pérez-

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Coello, M. S. Volatile Components and Key Odorants of Fennel (Foeniculum

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vulgare Mill.) and Thyme (Thymus vulgaris L.) Oil Extracts Obtained by

598

Simultaneous Distillation-Extraction and Supercritical Fluid Extraction. J. Agric.

599

Food Chem. 2005, 53, 5385-5389.

600 601

602



603

Page 30 of 46

FIGURE CAPTIONS

604 605

Figure 1. Germination of Brassica napus seeds during storage under moist conditions

606

for one (T1), two (T2), three (T3), and four (T4) days, respectively.

607 608

Figure 2. Principal component analyses including metabolites (A, B: carbohydrates,

609

organic acids, one cyclic polyol; C, D: amino acids; E, F: glucosinolates) of Brassica

610

napus seeds after storage under moist conditions for zero (T0), one (T1), two (T2),

611

three (T3), and four (T4) days, respectively. (A, C, E) Score plots with the percentage of

612

total variance explained by the principal components in parentheses, group median

613

scores as larger symbols, groups surrounded by convex hulls. (B, D, F) Loadings plots

614

with loadings as arrows and loadings axes at the top and right (bottom and left: score

615

axes as in score plots). For metabolite abbreviations, see Table 1; n = 9-10 (2 plates x 5

616

subsamples). Within the groups T1-T4, black dots in the symbols indicate subsamples

617

from plate A (symbols without dots: plate B).

618 619

Figure 3. Metabolic map showing changes in pool sizes of metabolites in Brassica

620

napus seeds after storage under moist conditions for one (T1), two (T2), three (T3), and

621

four (T4) days, compared to the common control group (T0 – zero days). Major pathway

622

intermediates are shown, whereas dashed arrows indicate that intermediates were

623

omitted. Metabolites, which were found in B. napus seeds, are given in black. Heatmap

624

stripes show the mean fold changes (mean metabolite pool size in group compared to

625

mean pool size in common control group) on a log2 scale using a color code (blue –


Page 31 of 46


626

lower metabolite pool size, yellow – higher pool size). Fold change thresholds of < 0.5

627

(log2 scale: < -1) and > 2 (log2 scale: > 1) are delineated in the color bar. Crosses in the

628

heatmap stripes indicate that fold changes were set to minimum or maximum,

629

respectively, as metabolites were found in at least half of the samples of one group but

630

were absent in the other group. Grey squares mean that no fold changes were

631

computed, as metabolites were found in less than half of the samples of one group and

632

were absent in the other group. Full names of metabolites are shown at the top right-

633

hand corner and in Table 1. Means of n = 9-10 (2 plates x 5 subsamples).

634 635

Figure 4. Principal component analysis of volatile compounds of Brassica napus oils

636

after storage of the corresponding seeds under moist conditions for zero (T0), one (T1),

637

two (T2), three (T3), and four (T4) days, respectively. (A) Score plot with the percentage

638

of total variance explained by the principal components in parentheses, group median

639

scores as larger symbols, groups surrounded by convex hulls. (B) Loadings plot with

640

loadings as arrows and loadings axes at the top and right (bottom and left: score axes

641

as in score plot). No loading is shown for PrOH, as it was too short for illustration. For

642

metabolite abbreviations, see Table 2. TAGs are given with their respective numbers

643

(see Figure S1); n = 10 (2 plates x 5 measurements over time). Within the groups, black

644

dots in the symbols indicate the oil pressed from plate A (symbols without dots: oil from

645

plate B). Roman numbers represent the day of measurement of the oil.

646 647

Figure 5. Sensory quality of virgin oil pressed from Brassica napus seeds after storage

648

of the seeds under moist conditions for zero (T0), one (T1), two (T2), three (T3), and



Page 32 of 46

649

four (T4) days, respectively, presented as radar charts. For each attribute, means

650

(replicate oils A/B) of the medians (four panellists) are given for each treatment group.

651

Full names of sensory attributes are given in the top-right corner. Off-flavor attributes

652

are highlighted with a grey area.


Page 33 of 46


TABLES Table 1. Metabolites Identified in Brassica napus Seeds analytical b platform

a

metabolite

retention c parameter

chemical class

abbreviation

name

carbohydrates

Frc

fructose

GC-MS

1860/1870

Glc

glucose

GC-MS

Suc

sucrose

Raf

organic acids

cyclic polyols

characteristic m/z

d

identification

GMD

std

217/277/364/335/307

x

x

1884/1902

319/229/343/305/160

x

x

GC-MS

2616

451/361/319/157/437

x

x

raffinose

GC-MS

3350

451/361/217/204/437

x

x

Glyc

glyceric acid

GC-MS

1324

189/307/205/133/292

x

x

Mal

malic acid

GC-MS

1484

245/335/307/217/233

x

x

Cit

citric acid

GC-MS

1811

375/211/183/257/273

x

x

Dehy-Asc

dehydroascorbic acid

GC-MS

1839

173/157/245/231/316

x

Ino

myo-inositol

GC-MS

2075

265/318/191/507/305

x

ASP

aspartic acid

UHPLC-FLD

2.9

min

x

GLU

glutamic acid

UHPLC-FLD

4.6

min

x

x

amino acids primary


e

idb


secondary

Page 34 of 46

ASN

asparagine

UHPLC-FLD

8.7

min

x

SER

serine

UHPLC-FLD

9.4

min

x

GLN

glutamine

UHPLC-FLD

11.0

min

x

HIS

histidine

UHPLC-FLD

11.5

min

x

GLY

glycine

UHPLC-FLD

12.3

min

x

THR

threonine

UHPLC-FLD

12.7

min

x

ARG

arginine

UHPLC-FLD

15.0

min

x

ALA

alanine

UHPLC-FLD

15.7

min

x

GABA

γ-aminobutyric acid

UHPLC-FLD

16.4

min

x

TYR

tyrosine

UHPLC-FLD

19.0

min

x

VAL

valine

UHPLC-FLD

23.4

min

x

MET

methionine

UHPLC-FLD

24.1

min

x

TRP

tryptophan

UHPLC-FLD

26.3

min

x

PHE

phenylalanine

UHPLC-FLD

27.1

min

x

ILE

isoleucine

UHPLC-FLD

27.5

min

x

LEU

leucine

UHPLC-FLD

29.2

min

x

LYS

lysine

UHPLC-FLD

30.3

min

x

PRO

proline

UHPLC-FLD

38.8

min

x

2ROH3but

progoitrin (2-R-2-hydroxy-3-butenyl)

HPLC-DAD

6.3

min

glucosinolates

aliphatic

x


Page 35 of 46


4MSOB

glucoraphanin (4-methylsulfinylbutyl)

HPLC-DAD

7.4

min

x

2SOH3but

epiprogoitrin (2-S-2-hydroxy-3-butenyl)

HPLC-DAD

7.5

min

x

1ME

glucoputranjivin (1-methylethyl)

HPLC-DAD

13.3

min

(x)

5MSOP

glucoalyssin (5-methylsulfinylpentyl)

HPLC-DAD

15.0

min

x

3but

gluconapin (3-butenyl)

HPLC-DAD

17.2

min

x

4pent

glucobrassicanapin (4-pentenyl)

HPLC-DAD

22.7

min

x

5MTP

glucoberteroin (5-methylthiopentyl)

HPLC-DAD

27.1

min

x

benzyl

2PE

gluconasturtiin (2-phenylethyl)

HPLC-DAD

27.0

min

x

indole

4OHI3M

HPLC-DAD

18.2

min

HPLC-DAD

24.2

min

HPLC-DAD

27.2

min

HPLC-DAD

29.6

min

4-hydroxyglucobrassicin

x

(4-hydroxy-indol-3-ylmethyl) glucobrassicin I3M

x

(indol-3-ylmethyl) 4-methoxyglucobrassicin 4MOI3M

x

(4-methoxyindol-3-ylmethyl) neoglucobrassicin 1MOI3M

x

(1-methoxyindol-3-ylmethyl)

a

Within each chemical class, metabolites are ordered according to their retention time. Names of organic acids are given in protonated form. For glucosinolates, common names are given with the side chains in parentheses. b GC-MS – gas chromatography coupled to mass spectrometry; UHPLC-FLD – ultra-high performance liquid chromatography coupled to fluorescence detection; HPLC-FLD – HPLC coupled to diode array detection. c For GC-MS data, Kováts retention indices are given, whereas retention times are given for the other analytical platforms. If one metabolite had more than one analyte (GC-MS), retention indices of both analytes are shown. d m/z – mass-to-charge ratio. Characteristic m/z values for each metabolite are shown.



Page 36 of 46

e

It is indicated whether metabolites were identified via comparison of retention parameters, UV spectra (for HPLC-DAD), and mass spectra (for GC-MS) with entries in the Golm metabolome database (GMD; GC-MS data only), an internal glucosinolate database (idb; HPLC-DAD data only), and/or authentic standards (std). Crosses indicate successful identification, whereas missing crosses indicate that the corresponding metabolite was not deposited in the database or no standard was available. Crosses in parentheses mean that the metabolite could only tentatively be identified, as not all parameters used for identification fit well.


Page 37 of 46


Table 2. Volatile Metabolites Identified in the Headspace of Brassica napus Virgin Oil

a

b

metabolite

RI

chemical class

abbreviation

name

alcohols

EtOH

ethanol

537

PrOH

propan-1-ol

aldehydes

characteristic m/z

c

identifid cation NIST

std

45/46

x

x

657

41/42/59/60

x

x

g

f

sensory quality

g

2MePrOH

2-methylpropan-1-ol

735

41/42/43/74

x

BuOH

butan-1-ol

772

41/43/55/56 (74)

x

x

3MeBuOH

3-methylbutan-1-ol

842

41/42/43/55/57/70 (88)

x

x

2MeBuOH 2.3BuOHI 2.3BuOHII HexOH HepOH 1Oc3OH

2-methylbutan-1-ol h butane-2,3-diol I h butane-2,3-diol II hexan-1-ol heptan-1-ol 1-octen-3-ol

844 961 972 976 1075 1078

41/55/56/57/70 (88) 45/57 (90) 45/57 (90) 27/29/31/39/41/42/43/55/56/69/84 (102) 41/42/43/55/56/57/69/70/83/103 (116) 43/57/72/85 (128)

x x

x x x

x x x

x x

EtCHO 2MePrCHO

acetaldehyde 2-methylpropanal

487 608

29/43/44 39/41//43/72

x x

x x

BuCHO

butanal

661

27/29/39/41/43/44/57/72

x

3MeBuCHO

3-methylbutanal

735

44/58/71/86

x

x

2MeBuCHO PeCHO HexCHO

2-methylbutanal pentanal hexanal

739 779 884

39/41//57/58/86 41/44/57/58/86 41/43/44/56/57/72/83 (100)

x x x

x x x

HepCHO

heptanal

990

41/42/43/44/55/57/70/71/81/86/96 (114)

x

x

OcCHO

octanal

1093

41/42/43/44/55/56/57/69/84 (128)

x

x

g

log2 fold e change

10

solvent

mushroom

42,48

10

g

moldy, sweet pungent, unpleasant, 10,48 cheese-like malty, cheese-like, flea10,42,49 bitten 42,49 malty 10 cheese-like, moldy 10,42,48,49 green, grass green, sweet, lemon, flower-like, fatty, 10,48 rancid green, lemon, 10,49 grass



isothiocyanates

NoCHO

nonanal

1199

41/43/44/55/56/57/68/69/70/81/82/98 (142)

2Octenal

2-octenal

1181

27/29/39/41/42/55/57/67/69/70/82/83 (126)

2PrITC

isopropylisothiocyanate

935

41/43/60/86/101

x

x

AITC

allylisothiocyanate

997

39/41/72/99

x

x

1ITCBu

1-isothiocyanatobutane

1037

29/41/56/57/72/115

(x)

2BuITC

isobutylisothiocyanate 4-isothiocyanato-1butene cyclopentylisothiocyana te

1064

27/29/39/41/43/57/72/73/115

x

1107

55/72/85/113

x

1215

39/41/67/68/69/127

x

2.3BuO2

butane-2,3-dione

681

43/86

2HeO

heptan-2-one

986

43/58/71/114

x

3Oc2O

3-octen-2-one

1158

41/43/55/97/111/126

x

MeICN 2MePrCN 3PCN 5HexCN

methylisocyanide 2-methylpropanenitrile 3-pentenenitrile 5-hexenenitrile 2propenylthioacetonitrile

596 746 908 1011

40/41 28/41/42/54/68/69 39/41/54/81 39/41/55/67/80/95

x x x x

1357

39/41/45/73/113

(x)

COS

carbonylsulfide

460

32/60

x

CH4S

methanethiol

489

45/47/48

x

DMS

dimethylsulfide

542

45/46/47/61/62

x

4ITC1Bu CycPeITC ketones

nitriles

2PrSNCS

organosulfur compounds

Page 38 of 46

x

x x

green, citrus, detergent, soap, 10,48,49 sweet fatty, nutty, 10,42,49 roasted

sulfur, garlic, 48 pungent sulfury, pungent, 48 green

x

(x)

48,49

buttery, caramel musty, spicy, blue 50 cheese 48 nutty

x

x

sulfur, 48 cookedcabbage moldy, cheese-like, fleabitten, cabbage, cooked 10,48,49 cauliflower


Page 39 of 46


terpenes

others

DMDS

dimethyldisulfide

813

45/46/47/61/64/79/94

x

x

MeTC

methylthiocyanate

857

45/46/58/72/73

x

x

DMTS

dimethyltrisulfide

1060

44/47/64/79/110/126/128

x

x

DMSO

dimethylsulfone

1200

15/79/94

x

αPin βPin Lim Cam

α-pinene β-pinene limonene camphor

954 1010 1063 1279

77/79/91/92/93/105/121/136 41/69/77/79/91/93/121/136 67/68/79/93/107/121/136 39/41/55/69/81/83/95/108/109/110/152

x x x x

Et2O 3MePe MeChl MeCycPe 3MeFu Tol XylI XylII Et4PeOate Styr Myr 4But

diethylether 3-methylpentane methylenechloride methylcyclopentane 3-methylfuran toluene h xyleneI h xyleneII ethyl 4-pentenoate styrene β-myrcene γ-butyrolactone

518 554 575 611 644 820 917 924 958 963 1024 1142

31/45/59/74 29/41/43/56/57/71/86 49/51/84/86 (85) 41/55/56/69/84 27/39/53/81/82 65/91/92 77/91/105/106 77/91/105/106 27/29/39/54/55/83 (128) 51/77/78/103/104 27/39/41/69/93 (136) 27/28/29/41/42/56/86

x x x x x x x x

4Val

γ-valerolactone

1177

41/56/85/100

x x x x

cabbage, sulfur, 48 ripened cheese 48 sulfur sulfur, cauliflower, 10,42,49 cabbage

10,48,49

citrus, spicy 52 camphor, rancid, oily

sweet, fruity

x x x

x x x x x x x

x

x

mushroom, earth10 moist

10

a

Within each chemical class, metabolites are ordered according to their retention time. Kováts retention indices.

b



Page 40 of 46

c

m/z – mass-to-charge ratio. Characteristic m/z values for each metabolite as shown in the NIST Chemistry WebBook. Molecular ions are indicated in bold if they were part of the characteristic mass spectra, otherwise molecular ions are parenthesized. d It is indicated whether metabolites were identified via comparison of mass spectra with those deposited in the NIST Chemistry WebBook and/or via comparison of retention indices and mass spectra with those of authentic standards (std). Crosses indicate successful identification, whereas missing crosses indicate that the corresponding metabolite was not deposited in the database or no standard was available. Crosses in parentheses mean that the metabolite could only tentatively be identified, as not all parameters used for identification fit well. e Mean fold changes (log2 scale) of metabolite pool sizes after storage of seeds under moist conditions for one (T1), two (T2), three (T3), and four (T4) days, compared to the common control (T0) group, given as heatmap stripes. The color code (blue - lower metabolite pool size, yellow higher pool size) is given at the end of the column. Fold change thresholds of < 0.5 (log2 scale: < -1) and > 2 (log2 scale: > 1) are indicated in the color bar. Crosses in the heatmap stripes indicate that fold changes were set to minimum or maximum, respectively, as metabolites were found in at least half of the samples of one group but were absent in the other group. Grey squares mean that no fold changes were computed, as metabolites were found in less than half of the samples of one group and were absent in the other group. Means of n = 10 (2 plates x 5 measurements over time). f Volatiles were described to be aroma-active by the specified references. g Equal RIs, because volatiles could chromatographically not be separated. Thus, fold changes were calculated based on the sum of both metabolites. h Roman numerals describe R-/S-isomers of the same metabolite.


Page 41 of 46


FIGURE GRAPHICS Figure 1



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Figure 2


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Figure 3



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Figure 4


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Figure 5

653



Page 46 of 46

TABLE OF CONTENTS GRAPHIC


Metabolic Changes during Storage of Brassica ... - ACS Publications

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