Assessment of anti-nutritional compounds and chemotaxonomic

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Bioactive Constituents, Metabolites, and Functions

Assessment of anti-nutritional compounds and chemotaxonomic relationships between Camelina sativa and its wild relatives Lisa Amyot, Tim McDowell, Sara L. Martin, Justin Renaud, Margaret Y. Gruber, and Abdelali Hannoufa J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.8b04724 • Publication Date (Web): 20 Dec 2018 Downloaded from http://pubs.acs.org on December 22, 2018

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Journal of Agricultural and Food Chemistry

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Assessment of anti-nutritional compounds and chemotaxonomic relationships between

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Camelina sativa and its wild relatives

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Lisa Amyot1, Tim McDowell1, Sara L. Martin2, Justin Renaud1, Margaret Y. Gruber3, and

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Abdelali Hannoufa1 *

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1Agriculture

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2

8

Ave., Ottawa, ON K1A 06C, Canada

9

3Agriculture

and Agri-Food Canada, 1391 Sandford Street, London, ON, N5V 4T3, Canada

Agriculture and Agri-Food Canada, Ottawa Research and Development Centre, 960 Carling

and Agri-Food Canada, Saskatoon Research Centre, 107 Science Place, Saskatoon,

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SK, S7N 0X2, Canada (retired)

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*To whom correspondence should be addressed; Email: [email protected]

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Phone: 1-519-953-6621. Fax: 1-519-457-3997

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ACS Paragon Plus Environment


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Abstract

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We compared the secondary metabolite composition in seeds of Camelina sativa and its wild

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relatives to identify potential germplasm with reduced levels of anti-nutritional compounds.

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Twenty Camelina accessions, from five different species, were analyzed by LC-MS and

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subjected to PCA which revealed that Camelina spp. separated into distinct chemotaxonomic

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groups. Three major glucosinolates (GS) were identified in our study, namely, 9-

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methylsulfinylonyl GS (GS9), 10-methylsulfinylonyl GS (GS10), and 11-methylsulfinylonyl GS

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(GS11). While there were differences in total GS levels, species-specific patterns for GS9 and

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GS11 were noted. Sinapine content ranged between 1.4 and 5.6 mg/g FW, with the lowest levels

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observed in C. laxa and C. sativa. Lignin levels were also lowest in C. sativa, with most

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accessions containing less than 6 mg/g FW. Our results show that wild Camelina spp. have

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distinct metabolomes and based on their levels of major anti-nutritionals, some could be

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incorporated into breeding programs with C. sativa.

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Keywords: Camelina, chemotaxonomy, glucosinolate, isothiocyanate, nitrile, sinapine, lignin

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Introduction

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Camelina sativa is an emerging oil seed that is valued for its high omega-3 fatty acid content and

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its ability to tolerate colder climates and other abiotic stresses 1-2. The seed meal remaining after

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oil processing is rich in protein, accounting for an average of 35-40% of the meal, and therefore

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has the potential for use as a supplement in animal feed

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when poultry were fed a diet consisting of 10% C. sativa meal, omega-3 fatty acid content in the

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eggs and meat was enhanced by 8- and 3-fold, respectively, without affecting bird performance.

3-4.

In fact, Cherian (2012) 4 noted that

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Despite its attributes, the use of C. sativa meal as a supplement in animal feed is still

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limited due to the presence of anti-nutritional compounds that can be toxic at high levels or that

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could affect meal palatability

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which are major anti-nutrients in Cruciferous species 7. Considerable efforts have been made

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over the years to reduce the levels of these compounds in rapeseed (Brassica napus). In their

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review, Khajali and Slominski (2012) 8 reported that glucosinolates in canola grade B. napus are

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approximately 1/12th of what they were originally, and that B. napus meal can now make up as

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much as 20% of poultry diets. In Brassicas, there is little natural variation in sinapine content 9,

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limiting the usefulness of conventional breeding in reducing sinapine levels. Instead, molecular-

63

based approaches involving downregulation of genes of the sinapine biosynthesis pathway have

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been pursued, which resulted in up to a 90% reduction in sinapine content 10.

5-6.

Of particular concern are glucosinolates, lignin, and sinapine,

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Sinapine is a choline ester of monomeric sinapic acid which arises from the

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phenylpropanoid pathway. Sinapine promotes seed germination by up-regulating UDP-

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glucotransferase genes involved in abscisic acid catabolism 11. While sinapic acid is considered a

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potent anti-oxidant with anti-microbial, anti-cancer, and anti-inflammatory activities

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imparts an unpleasant odour and taste on the meal that can affect its palatability and


12,

it also


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subsequently the quality of animal by-products 13. For example, when brown-shelled egg laying

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hens were fed a diet high in sinapine, their eggs developed a “fishy” taste and odour 8. Although

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sinapine content has been reported to be variable in Camelina spp., a number of studies have

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indicated that Camelina has lower levels compared to other Brassicaceae family members such

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as B. napus 6-7.

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Lignin is a collection of phenolic polymers that like sinapine arises from the

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phenylpropanoid pathway. Due to its indigestibility in monogastric animals and poor digestibility

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in ruminants, high lignin content lowers the energy value of animal feed and leads to reduced

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feed intake

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various protein sources for animal feed, demonstrated that Camelina seed meal had the lowest

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lignin content, as a percentage of dry matter, compared to meals from Brassica carinata, canola,

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linseed, and soybean 15. Colombini et al. (2014) 6 also showed that seed meal from Camelina had

82

a lower acid detergent lignin content and higher net energy levels compared to B. napus. While

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these studies are promising, there has not been extensive research into lignin levels of wild

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Camelina relatives.

14.

A recent study evaluating ruminal degradation and intestinal digestibility of

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Glucosinolates (GSs) are another class of anti-nutritional metabolites that reduce the

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palatability of meals derived from cruciferous seeds. Three major methionine-derived aliphatic

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GSs have been reported in Camelina: 9-methyl-sulfinyl-nonyl GS (GS9), 10-methyl-sulfinyl-

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decyl GS (GS10), and 11-methyl-sulfinyl-undecyl GS (GS11) 1. The bitterness attributed to these

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compounds is actually due to their hydrolysis products which can also be toxic at high levels,

90

affecting thyroid function and growth

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prior to the development of low-GS Brassica varieties (canola), they had been shown to

16.

GSs are of particular concern in poultry feed, where


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adversely affect egg production in laying hens as well as reduce the growth rate and increase

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mortality in broiler chickens fed on rapeseed meal 8.

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GSs are part of the plant defense system where upon wounding by pests or pathogens,

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they are hydrolyzed by myrosinase enzymes to produce bioactive compounds such as

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isothiocyanates and nitriles

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indicated that the effects on insect feeding and oviposition behavior may be determined more by

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the specific type of GS than by the total content of GSs 18. The same may also be true when it

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comes to evaluating toxicity of GSs in seed meal. In fact, some GSs have been reported to have

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health benefits. Interestingly the methyl-sulfinyl GSs in Camelina belong to the same class as

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glucoraphanin (4-methyl-sulfinyl-butyl GS), the precursor to the broccoli isothiocyanate,

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sulforaphane, which has been touted as having anti-cancer and anti-inflammatory properties

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Isothiocyanates with a methyl-sulfinyl-alk(en)yl side chain have been shown to be better

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inducers of the detoxifying glutathione S-transferases than those lacking an S-oxidized alk(en)yl

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group 20 .

17.

Studies comparing Brassica species with different GS profiles

19.

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The goal of the current study was to investigate natural variations in levels of sinapine,

107

lignin, and GSs in seeds of C. sativa and some of its wild relatives to determine if there is

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potential for introducing wild germplasm into breeding programs. Using a chemotaxonomy

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approach we were able to show that Camelina spp. have distinct metabolic profiles. Importantly

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we were also able to identify isothiocyanate and nitrile hydrolysis products of the major

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Camelina GS.

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Materials and Methods



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Seeds. Seeds of C. sativa, C. rumelica subsp. rumelica, C. microcarpa (cytotypes 2x, 4x,

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and 6x), C. laxa, C hispida subsp. hispida, and C. hispida subsp. grandiflora were generated at

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the Ottawa Research and Development Centre, Agriculture and Agri-Food Canada using seed

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from the North Central Regional Plant Introduction Station Ames, Iowa, USA (NCSPIS); the

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Leibniz Institute of Plant Genetics and Crop Plant Research, Gatersleben, Germany (IPK); the

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Plant Gene Resources of Canada, Saskatoon, Saskatchewan, Canada (PGRC); material collected

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from the Canadian Prairies; and several cultivars of C. sativa (Table 1). C. sativa 596, and

121

accessions from all other Camelina spp. required stratification at 4oC in the dark for two weeks.

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This was done in petri dishes on filter paper moistened with 2% potassium nitrate. C. sativa 596,

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C. rumelica, and C. microcarpa required an additional vernalization treatment once they reached

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the rosette stage. For vernalization, seedlings were transferred to 48 cell trays and grown under a

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16-hour photoperiod at 4oC for six weeks. The seeds were produced under a 16-hour photoperiod

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in a growth chamber set at 20oC with randomized individual position and re-randomization of

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position every two weeks. Stratification and vernalization treatments were planned so that plants

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could all be put in the growth chamber for seed production at the same time. Replicate number

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(parent plant) per accession ranged from 4-16 and the self-incompatible taxa were hand

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pollinated to induce seed set. Camelina spp. are referred to by their ID# in Table 1 and by text in

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the Results and Discussion section.

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Chemicals. All chemicals were purchased from Sigma-Aldrich (Oakville, Ontario),

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except for the sinapine standard which was kindly provided by Dr. Alfred Baumert from the

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Department of Secondary Metabolism, Leibniz Institute of Plant Biochemistry, Halle (Saale),

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Germany, and the HPLC-grade water and methanol (MeOH) which were purchased from Fisher


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Scientific (Markham, Ontario). Sigma-Aldrich standards for calibration curves included (−)-

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sinigrin hydrate (cat#00290) and lignin alkali (cat# 370959).

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HPLC Profiling. Metabolites were extracted from 0.05 g of ground, freeze-dried seed

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with 750 µL of 50% MeOH, 1.5% acetic acid (v/v) by mixing on a VWR multi-vortexer

140

(Mississauga, Ontario) at 2,500 rpm for 5 min. After a 5 min incubation on ice, the samples were

141

centrifuged (19,357 g, 5 min) and filtered through Acrodisc 0.22 µ 13 mm nylon filters (PALL

142

Industries, Mississauga, Ontario) into vials.

143

Filtered extracts were analyzed on an Agilent 1260 Series HPLC system (Agilent

144

Technologies, Mississauga, Ontario) equipped with a G4212B photodiode array detector,

145

G7112B Infinity II binary pump, G7129A autosampler, and a G1316A thermostat column

146

compartment set at 32oC. An HPLC-DAD gradient method, described below, was developed for

147

optimal separation of the major peaks from 4 µL of extract injected on a Poroshell 120 EC-C8

148

2.7µm 4.6 x 100 mm column fitted with a Poroshell 120 Fast Guard EC-C18 guard column

149

(Agilent) using a flow rate of 0.5 mL/min. The mobile phase consisted of water (A) and MeOH

150

(B), with both solvents containing 0.1% TFA. Elution was carried out over 30 min using the

151

following program which included conditioning: isocratic at 10% B for 1 min, then consecutive

152

increasing linear gradients from 10-35% B in 7 min, 35-75% B in 15.5 min, and 75-100% in 0.1

153

min, followed by isocratic 100% B for 2 min, linear gradient decrease to 10% B in 0.5 min, and

154

finally to isocratic 10% B for 2.5 min. Analytes were monitored at 229, 254, 332, and 360 nm.

155

Chromatographs from each channel were compared to determine whether there were species

156

specific patterns with respect to relative peaks heights and presence or absence of peaks. Peaks

157

corresponding to sinapine and three major GSs were identified by collecting fractions with a

158

G1364C fraction collector and analyzing them by LC-MS using the method described below.



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Sinapine was further confirmed by comparing retention times and absorption spectra to a

160

standard, and also by co-chromatography (spiking with a standard). Compound amounts were

161

calculated from the peak area using a calibration curve of the standard and then adjusting for

162

seed fresh weight. For GSs, a sinigrin standard was used as a surrogate.

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LC-MS. All MS data were acquired on a Q-Exactive™ Quadrupole Orbitrap mass

164

spectrometer (Thermo Scientific, MA, USA) coupled to an Agilent 1290 high-performance

165

liquid chromatography (HPLC) system. Samples were resolved on a Zorbax Eclipse Plus RRHD

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C18 (2.1 × 50 mm, 1.8 µm; Agilent Technologies, CA, USA) column maintained at 35 °C. The

167

mobile phases were: (A) water with 0.1% formic acid and (B) acetonitrile with 0.1% formic acid

168

(B), both Optima grade (Fisher Scientific, NJ, USA). The gradient consisted of 0% B for 30 secs

169

before increasing to 100% over 3 min, held at 100% for 2.5 min and reduced to 0% over 30

170

seconds. The following conditions were used for HESI: capillary temperature, 400 °C; sheath

171

gas, 17.00 units; auxiliary gas, 8.00 units; probe heater temperature, 450 °C; S-Lens RF level,

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45.00. The capillary voltages were 3.9kV and 4.0kV for positive and negative ionization mode

173

respectively. For metabolomic profiling, samples were analyzed in both positive and negative

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ionization at 140,000 resolution, a scan range of 100-1500 m/z, an automatic gain control (AGC)

175

target of 3×106, and a maximum injection time (IT) of 512 ms. For compound identification,

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representative samples were analyzed by data-dependent acquisition methods in both positive

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and negative ionization modes that involved a full MS scan at 17,500 resolution, an AGC target

178

of 3×106, and a max IT of 64 ms. The 12 highest intensity ions from the full scan (excluding

179

isotopes) were sequentially selected using a 1.2 m/z isolation window and re-analyzed at 17,500

180

resolution; AGC target, 1×106; max IT, 64 ms; normalized collision energy (NCE) 35; threshold

181

intensity 1.6×105; and dynamic exclusion of 10 s.


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9

Metabolomics Data Analysis. The .raw files were converted to mzML format using

182

21

183

ProteoWizard

and chromatogram alignment and deconvolution were performed with XCMS

184

22.

185

tolerance of 5, and a signal to noise threshold of 5. The “obiwarp” method 24 was used to correct

186

retention times with allowable deviation of 5s. Features with areas below the signal to noise

187

threshold were integrated using the “fillPeaks” function and remaining zeros were imputed with

188

two-thirds the minimum value on a per mass basis 25 prior to log transformation.

Features were detected using the “centwave” algorithm

23:

a noise level of 3×106, a ppm

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Principal component analysis (PCA) was performed using pareto scaling with the

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“FactoMineR” package in R version 3.5.0 (Platform: x86_64-w64-mingw32/x64 (64-bit)).

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Eigenvalues were calculated for dimensions 1-5 and the cumulative percent of variance was

192

calculated for each dimension. For dimensions 1 and 2, data was sorted according to eigenvalues

193

and the features showing the highest variance in either direction of each of the dimensions was

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recorded. Features were putatively identified by using the measured m/z values to determine

195

chemical formula and comparing experimental MS/MS fragmentation patterns with those

196

available in literature and online spectral databases 26.

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Lignin Extraction and Quantification. Cell wall residue (CWR) was prepared from

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0.05 g of seeds ground in liquid nitrogen. Proteins were removed by extracting with 1 mL of 100

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mM potassium phosphate buffer, pH 7.8, containing 0.5% TritonX-100 (v/v). After addition of

200

the buffer, samples were vortexed briefly and then put in an Eppendorf thermomixer (1,400 rpm,

201

5 min). Following mixing, the samples were incubated on ice for 5 min and centrifuged at 20,817

202

g for 5 min. The extraction was repeated 3 times and then the pellets were washed with 1 mL of

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80% MeOH by vortexing briefly and then incubating 1 hour at 80oC in a thermomixer set at

204

1,000 rpm. Samples were cooled on ice for 5 min before centrifuging for 5 min at 2,665 g. The



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80% MeOH wash was repeated twice but incubation at 80oC was reduced to 5 min for the

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subsequent washes. A final wash with 1 mL of 100% MeOH was carried out in the same fashion

207

as the additional 80% MeOH washes. Pellets were dried overnight in a thermomixer set at 80oC.

208

The thioglycolic acid (TGA) method was used to derivatize lignin in CWR to make it 27.

209

soluble under alkaline conditions

Briefly, the pellets from the above CWR preparation were

210

resuspend in a mixture of 750 µL deionized water, 250 µL hydrochloric acid, and 100 µL TGA.

211

After briefly vortexing, the samples were incubated at 80oC for 3 hr and then centrifuged at

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20,817 g for 5 min. The pellets were washed with 1 mL of deionized water by vortexing and then

213

centrifuging at 20,817 g for 5 min. Lignin was solubilized by incubating with of 1 mL sodium

214

hydroxide (1 M) for 12 hr at room temperature on a rotator set at a slow speed. Cell debris was

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removed by centrifugation (20,817 g, 10 min) and the supernatant containing derivatized lignin

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was transferred to a new 2 mL screw cap tube. Hydrochloric acid (200 µL) was added to the

217

supernatant and the tubes were incubated for 4 hr at 4oC. The supernatant was removed by

218

centrifugation 20,817 g, 5 min) and the pellets were redissolved in 1 mL sodium hydroxide (1

219

M). Residual debris was removed by filtering through an Acrodisc 0.22 µm 13 mm filter and the

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filtrate was diluted 20-fold with sodium hydroxide (1 M). Absorbance was measured at 280 nm

221

using a Thermo Scientific, Multiskan GO Microplate Spectrophotometer. Samples were run in

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triplicate and lignin was calculated from corrected absorbance values using a standard calibration

223

curve and adjusting for seed fresh weight.

224 225

Results and Discussion

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Camelina sativa seed is used primarily for its oil, but the meal is of potential value as an additive

227

in livestock feed. Phytochemicals make up approximately 10% of C. sativa seed meal and have


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the potential to contribute to animal and even human nutrition, but they also include compounds

229

that can be toxic at high levels or that affect the palatability of the meal 1. This study was aimed

230

at comparing the secondary metabolite composition of the seeds of C. sativa and its wild

231

relatives to identify potential germplasm that could be incorporated into breeding programs.

232

While components such as lignin affect the digestibility of seed meal, sinapine and GSs are of

233

particular concern since they are considered to be major anti-nutritionals of the Brassicaceae

234

family 7. Hence emphasis was placed on these three groups of metabolites.

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Camelina spp. separate into distinct chemotaxonomic groups. To determine whether

236

there are chemotaxonomic relationships between Camelina accessions, we performed principal

237

component analysis (PCA) on 1713 LC-MS positive mode features of seed extracts from 20

238

accessions. Our results revealed that the germplasm accessions from the five Camelina spp. in

239

our study separate into four distinct groups according to their seed metabolic composition

240

(Figure 1). The first component accounted for 32.5% of the variance and distinguished Groups 3

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and 4 from each other as well as from Groups 1 and 2. Component 2 accounted for an additional

242

14.7% of the variance and separated Groups 1 and 2 from each other as well as from Groups 3

243

and 4.

244

As expected, accessions within a species tended to group together; however, it should be

245

noted that C. laxa and both accessions of C. hispida have a closer chemotaxonomic relationship

246

to each other than to the other species. Regardless, only a few accessions of these two species

247

were available to us for analysis. These three taxa are all self-incompatible diploids (2x) with

248

similar morphological characteristics including relatively large (>2 cm) flowers and obovoid-

249

shaped pods28. Interestingly, the C. microcarpa accessions which make up Group 3 appear to

250

cluster according to their ploidy, with 2x being the most distinct. This clustering of cytotypes is



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12 251

even more prevalent when PCA is conducted with LC-MS negative mode data (Supplementary

252

Figure 1). Although it was closest to Group 3, C. microcarpa 2x was actually midway between

253

Groups 3 and 2, separating from them in Dimensions 1 and 2, respectively. While it is tempting

254

to suggest that this clustering is due to ploidy, geographical isolation cannot be ruled out given

255

that the 2x cytotype originated from France (but was grown in Canada) and the 4x and 6x

256

accessions originated from Canada.

257

Camelina spp. have distinct chromatographic fingerprints. In order to identify the

258

major metabolites in Camelina seed extracts, we developed an HPLC method for optimal peak

259

separation in four different UV channels. Our original goal was to fractionate a pooled seed

260

extract for downstream LC-MS-MS analysis, but it became clear that the HPLC chromatographs

261

could also be used for taxonomic classification as they revealed species-specific peak patterns.

262

Chromatographic fingerprinting has been reported as an identification and screening tool for

263

medicinal plants

264

screening tool for Camelina breeding.

29-30,

and based on our results described below, it could also be a useful

265

A comparison of relative peak areas and absence/presence of chromatographic peaks

266

provides a visual representation of species and sub-species trends as indicated by representative

267

chromatographs for accessions used in the metabolomics study (Supplementary Figure 2). In

268

some cases, all samples within a group showed the same pattern. For example all of the C. sativa

269

accessions from Group 1 had peaks 5 and 6 but lacked detectable levels of peaks 4 and 10, which

270

was a trend unique to this species (Supplementary Figure 2a). The presence of peak 10 was

271

characteristic of Group 2 C. rumelica subsp. rumelica (Supplementary Figure 2b), although

272

differences between accessions were noted. Prominence of peak 6 was characteristic for Group 3

273

C. microcarpa samples; however, the 2x cytotype was distinct from the 4x and 6x cytotypes,


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since it also contained peak 5 and detectable, albeit low levels of peaks 10 (Supplementary

275

Figure 2c). This is consistent with the PCA results shown in Figure 1 and Supplementary Figure

276

1 where C. microcarpa 2x separates from the other two cytotypes in Dimension 1. While peak 5

277

was prominent in all Group 4 species, C. hispida subsp. grandiflora was distinguished as being

278

the only one that contained peaks 7 and 8 (Supplementary Figure 2d).

279

Sinapine levels varied within Camelina accessions and between species. Sinapine is

280

an important anti-nutrient that accumulates in mature seeds of C. sativa 31 and has been shown to

281

be positively correlated with oil levels in B. napus and B. carinata breeding lines 9. As the most

282

abundant esterified phenolic compound in the Brassicaceae, sinapine was the major peak

283

observed in the HPLC chromatographs at 332 nm (Peak 2, Supplementary Figure 2). Sinapine

284

had a positive mass ion of 310.15244 and fragmented into daughter ions with mass-to charge

285

(m/z) ratios of 175, 207, and 251. In total, 186 samples were analyzed, which included four

286

additional accessions for C. sativa. Sinapine content ranged between 1.4 and 5.6 mg/g FW

287

(Figure 2). Given the unequal variances, a Kruskal-Wallis test was performed which revealed

288

statistically significant differences in sinapine content between species (Supplementary Table 1).

289

In general, C. laxa and C. sativa had the lowest sinapine levels, ranging between 1.7-2.6

290

mg/g FW and 1.6-3.8 mg/g FW, respectively. Our results for C. sativa (68 samples from 9

291

different accessions) are similar to those of Matthäus and Zubr (2000) 32 who reported a range of

292

1.7-4.2 mg/g for 10 different accessions. Some of the wild species in our study showed variations

293

in sinapine levels. For instance, biological replicates from C. hispida subsp. hispida had both the

294

lowest and highest sinapine levels. Furthermore, C. rumelica subsp. rumelica tended to have the

295

highest levels (the majority with 4.0-5.3 mg/g FW), but it also had accessions with as little as 1.7

296

and 1.9 mg/g FW. Russo and Reggiani (2017)

7

also noted high variability in sinapine content



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among 47 C. sativa accessions, with levels ranging between 1.09 to 4.75 mg/g DW. However,

298

they pointed out that sinapine levels were still lower than those of canola-grade B. napus and

299

other Brassicaceae. Matthäus and Zubr (2000) 32 also reported lower sinapine levels in C. sativa

300

compared to mustard and rapeseed which had sinapine contents of 7 and 13 mg/g, respectively.

301

Despite the variations in sinapine content, the percentage of peak area for sinapine at 332

302

nm was more consistent between species and within accessions. For example, all biological

303

replicates of C. sativa had values of 50% and lower; whereas C. hispida subsp. hispida and C.

304

hispida subsp. grandiflora biological replicates were all 54% or higher (Figure 2), indicating that

305

C. sativa has either more or higher levels of other compounds absorbing at 332 nm. This

306

observation is supported by the distinct species-specific chemotaxonomic profiles discussed

307

above (Figure 1) which are also visualized in the HPLC chromatographs (Supplementary Figure

308

2).

309

Camelina spp. have distinct and significantly different seed lignin content. Lignin is

310

another important anti-nutritional factor that contributes to the poor digestibility of C. sativa seed

311

meal in monogastric animals 4. Our results showed that there were statistically significant

312

differences in seed lignin content between Camelina spp. (Figure 3) as determined by one-way

313

ANOVA (F (23,173) = 69.56, p

Assessment of anti-nutritional compounds and chemotaxonomic

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