Development of an efficient glucosinolate extraction method
© The Author(s) 2017
Received: 18 August 2016
Accepted: 11 March 2017
Published: 21 March 2017
Glucosinolates, anionic sulfur rich secondary metabolites, have been extensively studied because of their occurrence in the agriculturally important brassicaceae and their impact on human and animal health. There is also increasing interest in the biofumigant properties of toxic glucosinolate hydrolysis products as a method to control agricultural pests. Evaluating biofumigation potential requires rapid and accurate quantification of glucosinolates, but current commonly used methods of extraction prior to analysis involve a number of time consuming and hazardous steps; this study aimed to develop an improved method for glucosinolate extraction.
Three methods previously used to extract glucosinolates from brassicaceae tissues, namely extraction in cold methanol, extraction in boiling methanol, and extraction in boiling water were compared across tissue type (root, stem leaf) and four brassicaceae species (B. juncea, S. alba, R. sativus, and E. sativa). Cold methanol extraction was shown to perform as well or better than all other tested methods for extraction of glucosinolates with the exception of glucoraphasatin in R. sativus shoots. It was also demonstrated that lyophilisation methods, routinely used during extraction to allow tissue disruption, can reduce final glucosinolate concentrations and that extracting from frozen wet tissue samples in cold 80% methanol is more effective.
We present a simplified method for extracting glucosinolates from plant tissues which does not require the use of a freeze drier or boiling methanol, and is therefore less hazardous, and more time and cost effective. The presented method has been shown to have comparable or improved glucosinolate extraction efficiency relative to the commonly used ISO method for major glucosinolates in the Brassicaceae species studied: sinigrin and gluconasturtiin in B. juncea; sinalbin, glucotropaeolin, and gluconasturtiin in S. alba; glucoraphenin and glucoraphasatin in R. sativus; and glucosatavin, glucoerucin and glucoraphanin in E. sativa.
Glucosinolates, B-thioglucoside N-hydroxysulfate derivatives, are secondary metabolites found in brassicaceae and related families . Over 120 glucosinolates, which differ in variable aglycone side chains derived from an alpha-amino acid, have been identified and classified into aliphatic, aromatic and indole glucosinolates [2, 3]. Due to their prevalence in cultivated vegetables, spices, oils and animal feed, glucosinolates and their hydrolysis products have been much studied in the context of their effects on human and animal nutrition [4, 5]. Glucosinolates and their breakdown products have also been a focus of studies in dietary prevention of disorders linked to oxidative stress such as cancer and gastric ulcers [2, 6, 7] and more recently, potential undesirable dietary effects such as genotoxicity of glucosinolate breakdown products in broccoli  and Pak Choi . The breakdown of glucosinolates has also been studied because of their potential use as agricultural pesticides in a technique known as biofumigation. In biofumigation a glucosinolate-rich crop is mulched into the field, releasing toxic secondary glucosinolate by-products, in order to reduce the incidence of pests, weeds and diseases in the following arable and horticultural crops [10–13].
Myrosinase, an enzyme found in brassicaceae and compartmentalised in cells in close proximity to glucosinolates, is responsible for hydrolysing glucosinolates upon plant tissue disruption. Accurate analysis of glucosinolates therefore requires inactivation of myrosinase prior to tissue disruption. This is achieved by first freezing then freeze drying the tissue which allows disruption by milling or grinding to occur in the absence of water (Fig. 1). Lyophilisation, or freeze drying, is used to remove water from glucosinolate-containing tissues while preventing myrosinase mediated glucosinolate hydrolysis through thermal inhibition. Publications on freeze drying plant tissue have focussed primarily on the production of heat or its implications in generating oxygen sensitive foodstuffs (e.g. space, military or extreme-sport foodstuffs and instant coffee) . To our knowledge, no study has yet examined the efficiency of freeze drying in maintaining glucosinolate concentrations. Freeze drying functions on the principle of sublimation: pressure is reduced below the triple point of water (6.12 mbar, 0.01 °C) at which point sublimation of ice from the sample occurs. The cooling effect of sublimation should be high enough to ensure the sample remains below 0 °C for the initial stage of freeze drying, thus minimizing enzyme-driven glucosinolate hydrolysis. Rapid sample loading and rapid initial pressure drop are also required to avoid sample defrosting before pressure is reduced below 6.12 mbar. Leaves have a high surface area to volume ratio and may defrost quickly, activating myrosinase and reducing final glucosinolate concentration. Despite the importance of the freeze drying process in glucosinolate extraction, many authors do not report details which are likely to affect final concentrations of glucosinolates (e.g. how samples are transported, temperature of the room, whether a heating/cold plate is used and time taken for the pressure to drop).
The most commonly used methods for extraction of glucosinolates from plant material are based on the ISO 9167-1 method [15; highlighted in grey in Fig. 1], which was designed for extraction of glucosinolates from B. napus seed and has been adapted to suit the needs of researchers examining glucosinolate profiles of other plant species and tissue types. Although freeze drying is not explicitly detailed in the ISO 9167-1 method, it is an implicit requirement in order to avoid myrosinase mediated glucosinolate hydrolysis during disruption of leaf, stem or root tissues. Once the plant tissue is prepared, the ISO 9167-1 extraction is carried out at 75 °C in 70% methanol for 10 min. Heating the sample is thought to be an essential step to denature myrosinase, thus preventing enzymatic hydrolysis of glucosinolates . Samples are subsequently desulfated by ion exchange chromatography on a DEAE Sephadex column to remove impurities. Desulfoglucosinolates are then separated and identified using HPLC with a reverse phase C18 column and a UV or MS detector. Hazards associated with boiling methanol  and the time required for extractions using this method have led researchers to seek alternatives. Replacing heated methanol with boiling water is reported to have comparable [18, 19], and in some cases better , extraction efficiencies. Although most glucosinolates are thermostable for the typical 10–30 min heating period, indole glucosinolates such as 4-hydroxy-glucobrassicin and 4-methoxyglucobrassicin have been reported to degrade quickly at temperatures below 100 °C . In addition, prior to 2002 the major glucosinolate in leaves of E. sativa, 4-mercaptobutyl glucosinolate, was missed because it self-dimerises via formation of disulphide linkages during extraction . A major challenge therefore to ensuring consistent and repeatable GSL analysis is to create extraction conditions in which myrosinase is inactive, and glucosinolates do not self-react or degrade. A single study, conducted exclusively on radish roots, has demonstrated that cold extraction in 80% methanol does not cause appreciable reduction in glucosinolate concentrations compared to more conventional heated extraction methods . However, myrosinase activity can vary dramatically  and whether this method is suitable for extraction of glucosinolates from other glucosinolate containing plants has not previously been assessed.
A desulfation step is often carried out post extraction to purify desulfoglucosinolates and improve accuracy and identification from HPLC. However, the desulfation reaction of glucosinolates can be affected by feedback inhibition of the enzyme which causes incomplete desulfation of glucosinolates . In addition, rhamnopyranosyloxy-benzyl glucosinolates extracted from M. oleifera have been shown to be completely converted and degraded by the desulfation purification step . Due to these drawbacks, and the additional time and potential error extra steps can introduce, some authors have skipped the purification and desulfation steps entirely [19, 26, 27] (Fig. 1).
How do lyophilisation conditions affect glucosinolate concentrations?
Is lyophilisation a necessary step for glucosinolate extraction from green tissues?
Do extractions in hot methanol, cold methanol and boiling water yield comparable glucosinolate concentrations across a range of brassicaceae species and tissue types?
How do desulfation time and enzyme concentration affect final glucosinolate concentrations?
Is desulfation a necessary step for glucosinolate extraction from green tissue?
B. napus used in the freeze drying tests were grown in 1 L pots filled with Terra-green in a controlled temperature glasshouse (regulated from 17.6 to 27.7 °C). At 3–4 weeks post germination, leaves were removed and halved down the limits of the midrib, excluding the midrib from the final sample. Leaf halves were immediately frozen in liquid nitrogen and stored at −80 °C for a maximum of 1 week.
B. juncea (cv. ISCI99), R. sativus (cv. Bento), S. alba (cv. Ida Gold) and E. sativa (cv. Nemat) plants were grown by Barworth agriculture ltd. in a sandy loam soil dominated fields (coordinates: 53.000371, −0.290404) from 31/07/2014 to 25/09/2014. Total stem and total leaves were cut from flowering plants and immediately frozen in liquid nitrogen; root samples were gently washed and dried before freezing in liquid nitrogen. Samples were stored at −80 °C for a maximum of 2 months.
Freeze drier characteristics
Room temp (°C)
Time to 5 mbar (s)
Lowest pressure (mbar)
Freezer temperature (°C)
Lyotrap, LTE scientific ltd.
Thermo, Heto Powerdry LL3000
Freeze dried plant tissue was homogenised to a roughly ground powder (approximately 0.1 cm particle size) using a grinder (Lloytron, E5601BK) Homogenised ground samples were milled at a frequency of 20 Hz for 10 min (Retch, MM400) with 2 steel ball bearings to a fine powder (particle diameter <0.1 mm). Samples were then sealed and stored at 20 °C for up to 9 months.
- (ii)Frozen fresh B. napus leaf halves (experiment 2, Table 2) were placed in 2 ml eppendorf vials and stored at −20 °C. 1.755 ml of 80% methanol precooled at −20 °C, 25 µl of 5 mM sinigrin and 2 small ball bearings were added. Samples were milled for 10 min at frequency 20 Hz (TissueLyser II, Qiagen). Final concentrations of methanol were estimated by incorporating average leaf moisture content of fresh B. napus leaves according to Eq. (1). Final concentration of methanol ranged from 79.3 to 79.9% and leaf moisture content accounted for <1% of final liquid volume.Table 2
Summary of methods used
Freeze drying/tissue disruption
1—Effect of freeze drier on GSL concentration
FD-A or FD-B/mill
0.3 U/ml for 24 h
ISO 9167-1 method
2—Comparison of GSL extraction from freeze dried tissue with extraction from wet tissue
FD-A or −20 °C methanol
0.3 U/ml for 24 H
ISO 9167-1 method
3—Comparison of extraction methods
Leaves, stems, roots
0.3 U/ml for 24 H
ISO 9167-1 method
4—Comparison of desulfation/purification methods
Leaves, stems, roots
0.3 U/ml for 12, 24, 48 h, and 5 U/ml for 16 h or filtration
ISO 9167-1 method for desulfoGSL,
Herzallah and Holly method for intact GSLs
Extractions were carried out in one of three ways (Fig. 1). In each case 50 µl of a 5 mM gluctropaeolin (for B. juncea samples) or 20 mM sinigrin (for all other samples) internal standard was added.
Hot methanol extraction (based on the ISO 9167-1 method)
0.1 g of plant material was preheated at 75 °C for 3 min in a 20 ml falcon tube. 4.95 ml of 70:30 methanol:water, preheated to 75 °C and the internal standard was added. The sample was incubated at 75 °C for 10 min, and manually shaken every 2 min. The sample was then centrifuged at 4000 rpm (Jouan, model:B 3.11) for 10 min. Supernatent was stored at −20 °C or desulfated directly.
Cold methanol extraction (Ishida et al. )
5 ml of 80:20 methanol:water at 20 °C was added to 0.1 g plant tissue and the internal standard was added. The sample was shaken and left to stand for 30 min at room temperature. The sample was then mixed at 70 rpm with a platform rocker for a further 30 min (Bibby, STR6) before centrifugation at 4000 rpm (Jouan, model:B 3.11) for 10 min. Supernatent was then filtered through a 0.22 µm syringe filter (Millex GP) for direct injection on HPLC, or unfiltered if applied to Sephadex column in a purification step.
Boiling water extraction (adapted from Herzallah and Holley )
25 ml of boiling water was added to 0.1 g of freeze dried and milled plant tissue in a 150 ml erlenmeyer flask and the internal standard was added. Sample was heated at 100 °C and stirred with a magnetic stirrer hot plate for 10 min. Sample was heated for a further 4 h at 70 °C before centrifugation at 4000 rpm (Jouan, model:B 3.11) for 10 min. Sample was topped up to 20 ml with deionised water.
Purification and determination of activity of sulfatase
Sulfatase from Helix pomatia type H-1 (Sigma, S9626) was purified according to Wathalet et al. . 25 mg of sulfatase was added to 1 ml 40% ethanol and centrifuged at 8000 rmp for 1 min (eppendorf centrifuge, 54,151). The supernatant was transferred to a fresh 2 ml eppendorf tube, 1 ml of pure ethanol was added to precipitate the sulfatase before being centrifuged at 8 krmp for 1 min. The supernatant was discarded and the sulfatase pellet air dried and redissolved in 2 ml of water.
The activity for Sulfatase from Helix pomatia type H-1 (Sigma, S9626) given by the supplier is determined by desulfation of p-nitrocatechol sulfate and is an order of magnitude higher than the activity measured for desulfation of sinigrin using this method.
Desulfation of glucosinolates
As per the ISO 9067-1 method, columns were prepared with 0.5 ml Sephadex slurry (2 g DEAE Sephadex beads in 30 ml 2 M acetic acid.) and activated with 2 ml imizadole formate (6 M). Columns were washed twice with 1 ml water. The column was washed twice with 1 ml 20 mM sodium acetate (pH 4.0) and 75 µl of purified sulfatase was added (5 or 0.3 U/ml). Columns were incubated at room temperature for either 12, 24 or 48 h before elution of desulfoglucosinolates with two 1 ml volumes of water. For the reduction of disulphide linkages, from dimerized desulfoglucosatavin in E. sativa extracts 3 g TCEP (Tris(2-carboxyethyl)phosphine hydrochloride powder Sigma, C4706) was added to 1 ml of desulfated extract. Desulfoglucosinolates were stored at −20 °C before high performance liquid chromatography analysis (Additional file 1).
For the high sulfatase treatment, between 0.5 and 1 ml of sample was added due to insufficient sample volume remaining.
A Waters 600E system controller attached to a Waters 717 autosampler, Waters 996 photodiode array detector and SphereClone 5µ ODS(2) column (Phenomonex) were used for separation and detection of desulfo and intact glucosinolates.
HPLC analysis of desulfoglucosinolates—adapted from ISO 9167-1
Mobile phase conditions for separation of desulfoglucosinolates
% Solution A
% Solution B
Solution A: 100% diH2O
Solution B: 70:30, diH2O:acetonitrile
Desulfoglucosinolates were quantified using 229 nm wavelength within the UV spectrum. The HPLC PDA detector allowed a full spectrum analysis from 180 to 800 nm, allowing comparative UV–visible spectra analysis, which aided in identifying unknown glucosinolates. Through standard injections and HPLC–MS identification we were able to confirm the id’s of these reported glucosinolates. Desulfated purified standards: sinigrin (sigma aldrich), glucotropaeolin, glucoraphenin, glucoraphanin, glucerucin, glucobrassicin, gluconasturtiin, sinalbin, progoitrin and glucoiberin (phytoplan).
Source: Standard electrospray (flow split 1/10 from LC)
Nebulizer: 2.0 bar
Dry gas: 6.0 L/min
Dry gas heater: 25 °C
Capillary voltage: 3500 V
Ion polarity: positive
Spectra rate: 1 Hz
HPLC analysis of intact glucosinolates—adapted from Herzallah and Holly 
Solution A: 100% TBA (0.02 M)
Solution B: 70:30, TBA (0.02 M):acetonitrile
Glucosinolates were quantified using the chromatogram from 229 nm and standard curves were constructed using pure sinigrin (sigma aldrich), glucotropaeolin, glucoraphenin, glucoraphanin, glucerucin, glucobrassicin, gluconasturtiin, sinalbin, progoitrin and glucoiberin (phytoplan).
In the case of glucoraphasatin in R. sativus leaves and glucotropaeolin in B. juncea minor alterations were made to avoid peaks co-eluting. The mobile phase programme for R. sativus leaves was 100% A for 5 min, followed by a 35 min linear gradient to 66% B followed by a 5 min linear gradient to 100% B followed by a 5 min linear gradient to 100% A. For B. juncea leaves, an isocratic 85:15, TBA (0.02 M):acetonitrile mobile phase for 70 min was used.
Determination of myrosinase activity
Activity of pure myrosinase was tested in water and 80% methanol solutions containing 0.25 mM sinigrin and 0.1 mM ascorbic acid, a myrosinase cofactor . Myrosinase was added at t = 0 and absorbance of sinigrin at 229 nm was measured over the course of an hour. Activity was measured at room temperature (25 °C).
Determination of glucosinolate thermostability
A 50 µl of 10 mM sinigrin, 10 mM glucotropaeolin, 10 mM glucobrassicin solution was added to 0.95 ml water or 70% methanol preheated to 100 or 75 °C respectively and sealed in 1.5 ml eppendorf tubes. Samples were maintained at either 100 or 75 °C for 5, 10, 30 and 60 min and intact glucosinolate concentrations analysed with HPLC following the adapted Herzallah and Holly method .
Calculation of glucosinolate content
Paired two tailed t test analysis were carried out on total B. napus glucosinolate content per leaf half in experiments 1 and 2 with Microsoft excel (Table 2). For determination of significance of effect of method on final glucosinolate content estimates in experiments 3 and 4 (Table 2), repeat measure ANOVA analyses were carried out for each glucosinolate with R statistical software package (version 3.3.1).
Results and discussion
Modifications to the ISO9167-1 method (specifically created for the extraction and analysis of glucosinolates from oil rape seed samples) are required for analysis of plant green tissues (leaves, stems and roots). A number of prior-to-analysis steps, such as sampling in the field, cleaning (if required), freezing, crushing, storage or/and shipping and reduction of sample amount have been discussed by Wathelet et al.  and are not revisited here. These preliminary steps are followed by lyophilisation, or freeze drying, to remove water from glucosinolate containing tissues while preventing myrosinase mediated glucosinolate hydrolysis through thermal inhibition. This process allows subsequent tissue disruption without risking glucosinolate degradation.
Freeze drier characteristics
Room temp (°C)
Time to 5 mbar (s)
Lowest pressure (mbar)
Freezer temperature (°C)
These results underline the need for a more substantive study to assess optimal conditions for freeze drying plant tissues for glucosinolate analysis. It is clear that differences in freeze drying can introduce significant variability in retained glucosinolate concentrations (Fig. 2a).
A cold methanol extraction method may be sufficient to (1) inactivate myrosinase and (2) efficiently extract glucosinolates, precluding the need for the lyophilisation step altogether. We tested this by comparing glucosinolates extracted from one half of a B. napus leaf in 80% methanol without freeze drying against glucosinolates extracted from the other half, first dried in freeze drier A and then extracted using the cold methanol extraction method.
Glucosinolates examined in this study
Species, tissue type
B. juncea L, S, R
R. sativus L, S, R
E. sativa L, S, R
E. sativa L, S, R
Glucoraphasatin or hydroxyglucoerucin
R. sativus L, S, R
E. sativa S, R
S. alba, R
S. alba L, S, R
S. alba L, S, R
B. juncea R
S. alba R
S. alba R
No glucosinolates were detected in a subset of samples extracted in cold water indicating the presence of active myrosinase leading to their degradation (data not shown). However, the cold methanol extraction did not significantly affect the concentration of the internal standard relative to the boiling methanol method (data not shown), providing additional evidence that myrosinase is inactivated in 80% methanol without heating (Fig. 5).
These data demonstrate that 80% cold methanol can be used instead of boiling methanol to extract glucosinolates across a broad spectrum of brassicaceae species and tissue types. With the exception of glucoraphasatin in R. sativus shoots, replacing hot 70% methanol with cold 80% methanol did not significantly reduce glucosinolate concentrations, yet marginally increased recovery of sinalbin in S. alba and sinigrin in B. juncea. It is advised, due to reduction in steps and hazard as well as improved or comparable glucosinolate recovery, that a cold methanol extraction is used instead of a boiling methanol extraction for most glucosinolate containing green tissues.
Shorter desulfation times and lower sulfatase concentrations resulted in underestimation of the concentrations of glucoraphenin from R. sativus, glucoraphanin and glucosatavin from E. sativa, sinigrin from B. juncea, and sinalbin from S. alba and an overestimation of the concentrations of glucoraphasatin in R. sativus roots (Fig. 9). The overnight (12–24 h) incubation with 0.3 U/ml sulfatase solution yields inaccurate results for most major glucosinolates examined in this study. The ISO9167-1 method suggests that a diluted purified sulfatase solution with an activity exceeding 0.05 U/ml should be used, which is shown to be insufficient for glucosinolate analysis from plant samples and conditions examined in this study (Fig. 9). Instead, if a desulfation step is carried out, use of a higher concentration of purified sulfatase (in this case, 5 U/ml) is advised.
In all E. sativa leaf samples tested, recovery of monomeric desulfo-glucosatavin decreased and recovery of dimeric desulfo-glucosatavin increased between 24 and 48 h. Bennet et al.  previously hypothesised that dimeric glucosatavin is unlikely to be found in vivo and is probably an artefact of the extraction process. We can confirm that glucosatavin forms dimers as a result of the desulfation step of the extraction and that without carrying this step out and instead quantifying intact glucosinolates, no dimeric glucosatavin was detected in these samples.
Given that glucoraphenin concentration estimates are lower from methods employing a desulfation step, and that this step is also responsible for the dimerization of glucosatavin, analysis of intact glucosinolates is preferable in most instances. It is out of the scope of this study to compare or improve separation and detection methods but it should be noted that major glucosinolates in this study were accurately measured by a HPLC–UV method adapted from Herzallah and Holley . For examination of low abundance glucosinolates, and to avoid any potential inaccuracies due to contamination it is advised that an alternative HPLC method such as those suggested in Lee et al. or Forster et al. be used instead [26, 32].
Suggested method for glucosinolate extraction
Freeze samples loosely wrapped in foil in liquid nitrogen and store at −80 °C. Transport samples to freeze drier in dry ice. Rapidly load samples onto a cool plate in freeze drier and ensure the pressure drops to below 5 mbar in under 2 min. Mill samples once dried and store in airtight containers in the dark.
Freeze 50 mg samples in liquid nitrogen in 2 ml eppendorf tubes and store at −80 °C (for larger samples use larger tubes). Add a volume of 80% methanol precooled to −20 °C ensuring that final methanol concentration remains above 78% according to Eq. (1) in materials and methods. Add an appropriate volume of internal standard sinigrin or glucotropaeolin (e.g. 100 µM final concentration). Disrupt tissue by adding 2 small ball bearings and agitating with a tissue lyser (e.g. tissuelyserII, Qiagen) for 10 min at 20 rev/s. Alternatively use a plastic pestle to thoroughly grind the sample taking care that to keep the media below 0 °C. Continue directly to 2b.
For freeze dried tissue (1a). To 0.1 g tissue, add 5 ml of 80% methanol and 50 µL of 20 mM sinigrin solution. Then
Shake sample once and leave to stand for 30 min. Shake sample for a further 30 min (70 rev/s). Centrifuge at 4000 rpm and transfer supernatant to a fresh tube.
If desulfation is required, a high concentration sulfatase solution should be prepared by dissolving 15–25 mg sulfatase in 1 ml 40% ethanol and centrifuge at 8000 rmp for 1 min. Transfer supernatant to a fresh 2 ml eppendorf tube and add 1 ml of pure ethanol to precipitate the sulfatase and centrifuge at 8000 rpm for 1 min. Discard the supernatant and air dry the pellet before re-dissolving in 2 ml of water. Proceed with desulfation according to ISO9167-1 method.
In this study we compared different methods for extracting and purifying glucosinolates from B. napus, B. junea, S. alba, E. sativa and R. sativus green tissues to highlight unnecessary or hazardous steps. We have presented a simplified method for extracting glucosinolates from plant tissues which does not require the use of a freeze drier or boiling methanol, and is therefore less hazardous, and more time and cost effective. The presented method has been shown to have comparable or improved glucosinolate extraction efficiency relative to the commonly used ISO method for major glucosinolates in the Brassicaceae species studied: sinigrin and gluconasturtiin in B. juncea; sinalbin, glucotropaeolin, and gluconasturtiin in S. alba; glucoraphenin and glucoraphasatin (roots but not shoots) in R. sativus; and glucosatavin, glucoerucin and glucoraphanin in E. sativa.
TDA, KR, and SEH organized the project. VK carried out sample preparation and glucosinolate extractions on B. napus leaves. TDA performed all other experiments, analyzed the data, and wrote the paper; SEH, IB and KR reviewed and edited the manuscript. All authors read and approved the final manuscript.
We are grateful to Catherine Lilley, (University of Leeds), Peter Urwin (University of Leeds), Andy Barker (Barworth Agriculture ltd.) and Helen Barker (Barworth Agriculture ltd.) for collection of leaf, stem and root samples used in this study. We would also like to thank Thomas Hartley (University of York) for statistical advice. We would also like to express our gratitude to the BBSRC for funding this study. Finally, we would like to thank the reviewers for their time and effort in reviewing this manuscript.
The authors declare that they have no competing interests.
Availability of data and materials
Material used in this study is stored at the University of York and is available on request. Datasets analysed in this study are available from the corresponding author on request.
This work was supported by UK Biotechnology and Biology Sciences Research Council (BB/L002124/1) and (BB/K020463/1).
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- Clarke DB. Glucosinolates, structures and analysis in food. Anal Methods. 2010;2(4):310–25.View ArticleGoogle Scholar
- Fahey JW, Zalcmann AT, Talalay P. The chemical diversity and distribution of glucosinolates and isothiocyanates among plants. Phytochemistry. 2001;56(1):1–5.View ArticleGoogle Scholar
- Mithen RF, Dekker M, Verkerk R, Rabot S, Johnson IT. The nutritional significance, biosynthesis and bioavailability of glucosinolates in human foods. J Sci Food Agric. 2000;80(7):967–84.View ArticleGoogle Scholar
- VanEtten CH, Daxenbichler ME, Wolff IA. Natural glucosinolates (thioglucosides) in foods and feeds. J Agric Food Chem. 1969;17(3):483–91.View ArticleGoogle Scholar
- Cartea ME, Velasco P. Glucosinolates in Brassica foods: bioavailability in food and significance for human health. Phytochem Rev. 2008;7(2):213–29.View ArticleGoogle Scholar
- Talalay P, Fahey JW. Phytochemicals from cruciferous plants protect against cancer by modulating carcinogen metabolism. J Nutr. 2001;131(11):3027S–33S.PubMedGoogle Scholar
- Shapiro TA, Fahey JW, Wade KL, Stephenson KK, Talalay P. Chemoprotective glucosinolates and isothiocyanates of broccoli sprouts metabolism and excretion in humans. Cancer Epidemiol Biomark Prev. 2001;10(5):501–8.Google Scholar
- Latté KP, Appel KE, Lampen A. Health benefits and possible risks of broccoli—an overview. Food Chem Toxicol. 2011;49(12):3287–309.View ArticlePubMedGoogle Scholar
- Wiesner M, Schreiner M, Glatt H. High mutagenic activity of juice from pak choi (Brassica rapa ssp. chinensis) sprouts due to its content of 1-methoxy-3-indolylmethyl glucosinolate, and its enhancement by elicitation with methyl jasmonate. Food Chem Toxicol. 2014;31(67):10–6.View ArticleGoogle Scholar
- Ngala BM, Haydock PP, Woods S, Back MA. Biofumigation with Brassica juncea, Raphanus sativus and Eruca sativa for the management of field populations of the potato cyst nematode Globodera pallida. Pest Manag Sci. 2015;71(5):759–69.View ArticlePubMedGoogle Scholar
- Lord JS, Lazzeri L, Atkinson HJ, Urwin PE. Biofumigation for control of pale potato cyst nematodes: activity of Brassica leaf extracts and green manures on Globodera pallida in vitro and in soil. J Agric Food Chem. 2011;59(14):7882–90.View ArticlePubMedGoogle Scholar
- Mattner SW, Porter IJ, Gounder RK, Shanks AL, Wren DJ, Allen D. Factors that impact on the ability of biofumigants to suppress fungal pathogens and weeds of strawberry. Crop Prot. 2008;27(8):1165–73.View ArticleGoogle Scholar
- Bellostas N, Kachlicki P, Sørensen JC, Sørensen H. Glucosinolate profiling of seeds and sprouts of B. oleracea varieties used for food. Sci Hortic. 2007;114(4):234–42.View ArticleGoogle Scholar
- Ratti C. Hot air and freeze-drying of high-value foods: a review. J Food Eng. 2001;49(4):311–9.View ArticleGoogle Scholar
- ISO 9167-1, 1992 NA 057-05-05 AA—Joint committee of DIN and DGF for the analysis of fats, oils and products thereof, related and primary products. (2012): rapeseed—determination of glucosinolate content—part 1: method using high-performance liquid chromatography (ISO 9167–1:1992/DAM 1:2012), German version EN ISO 9167-1:1995/prA1: 2012.).Google Scholar
- Ares AM, Nozal MJ, Bernal JL, Bernal J. Optimized extraction, separation and quantification of twelve intact glucosinolates in broccoli leaves. Food Chem. 2014;1(152):66–74.View ArticleGoogle Scholar
- Church AS, Witting MD. Laboratory testing in ethanol, methanol, ethylene glycol, and isopropanol toxicities. J Emerg Med. 1997;15(5):687–92.View ArticlePubMedGoogle Scholar
- Rangkadilok N, Nicolas ME, Bennett RN, Premier RR, Eagling DR, Taylor PW. Determination of sinigrin and glucoraphanin in Brassica species using a simple extraction method combined with ion-pair HPLC analysis. Sci Hortic. 2002;96(1):27–41.View ArticleGoogle Scholar
- Herzallah S, Holley R. Determination of sinigrin, sinalbin, allyl-and benzyl isothiocyanates by RP-HPLC in mustard powder extracts. LWT Food Sci Technol. 2012;47(2):293–9.View ArticleGoogle Scholar
- Stoin DF, Dogaru RD. Researches regarding the isolation, purification and analysis of sinigrin glucosinolate from Brassica nigra and Armoracia rusticana. Bull USAMV-CN. 2007;63:77–82.Google Scholar
- Oerlemans K, Barrett DM, Suades CB, Verkerk R, Dekker M. Thermal degradation of glucosinolates in red cabbage. Food Chem. 2006;95(1):19–29.View ArticleGoogle Scholar
- Bennett RN, Mellon FA, Botting NP, Eagles J, Rosa EA, Williamson G. Identification of the major glucosinolate (4-mercaptobutyl glucosinolate) in leaves of Eruca sativa L. (salad rocket). Phytochemistry. 2002;61(1):25–30.View ArticlePubMedGoogle Scholar
- Ishida M, Kakizaki T, Ohara T, Morimitsu Y. Development of a simple and rapid extraction method of glucosinolates from radish roots. Breed Sci. 2011;61(2):208–11.View ArticleGoogle Scholar
- Piekarska A, Kusznierewicz B, Meller M, Dziedziul K, Namieśnik J, Bartoszek A. Myrosinase activity in different plant samples; optimisation of measurement conditions for spectrophotometric and pH-stat methods. Ind Crops Prod. 2013;31(50):58–67.View ArticleGoogle Scholar
- Wathelet JP, Mabon N, Marlier M. Determination of glucosinolates in rapeseed improvement of the official HPLC ISO method (precision and speed). In: Proceedings of the 10th international rapeseed congress. Gosford: The Regional Institute Ltd; 1999 p. 185.Google Scholar
- Förster N, Ulrichs C, Schreiner M, Müller CT, Mewis I. Development of a reliable extraction and quantification method for glucosinolates in Moringa oleifera. Food Chem. 2015;1(166):456–64.View ArticleGoogle Scholar
- Troyer JK, Stephenson KK, Fahey JW. Analysis of glucosinolates from broccoli and other cruciferous vegetables by hydrophilic interaction liquid chromatography. J Chromatogr A. 2001;919(2):299–304.View ArticlePubMedGoogle Scholar
- Wathelet JP, Iori R, Leoni O, Rollin P, Quinsac A, Palmieri S. Guidelines for glucosinolate analysis in green tissues used for biofumigation. Agroindustria. 2004;3(3):257–66.Google Scholar
- Karathanos VT, Anglea SA, Karel M. Structural collapse of plant materials during freeze-drying. J Therm Anal Calorim. 1996;47(5):1451–61.View ArticleGoogle Scholar
- Burmeister WP, Cottaz S, Rollin P, Vasella A, Henrissat B. High resolution X-ray crystallography shows that ascorbate is a cofactor for myrosinase and substitutes for the function of the catalytic base. J Biol Chem. 2000;275(50):39385–93.View ArticlePubMedGoogle Scholar
- Hennig K, Verkerk R, Bonnema G, Dekker M. Pitfalls in the desulphation of glucosinolates in a high-throughput assay. Food Chem. 2012;134(4):2355–61.View ArticlePubMedGoogle Scholar
- Lee JG, Bonnema G, Zhang N, Kwak JH, de Vos RC, Beekwilder J. Evaluation of glucosinolate variation in a collection of turnip (Brassica rapa) germplasm by the analysis of intact and desulfo glucosinolates. J Agric Food Chem. 2013;61(16):3984–93.View ArticlePubMedGoogle Scholar