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Determination of ergosterol in canola (Brassica napus L.) by liquid chromatography
Journal of Stored Products Research 39 (2003) 185–191
Determination of ergosterol in canola (Brassica napus L.) by liquid chromatography D. Abramson*, D.M. Smith Cereal Research Centre, Agriculture and Agri-Food Canada, 195 Dafoe Road, Winnipeg, Man., Canada R3T 2M9 Accepted 22 October 2001
Abstract A sensitive and precise method is described to assay the oilseed canola (Brassica napus L.) for ergosterol, a fungal metabolite indicating spoilage. The ground seed is refluxed in methanol, and the methanol extract is saponified with potassium hydroxide. After addition of water, the mixture is partitioned into n-hexane. The n-hexane extract is dried, reconstituted, and applied to a silica solid-phase extraction cartridge, which is then washed with carbon tetrachloride, and eluted with acetone. The acetone eluate is acetylated, and the ergosterol determined as the acetate by liquid chromatography using a reverse-phase column and absorbency detection at 282 nm. Acetylation is necessary since an unidentified constituent of the matrix cochromatographs with free ergosterol, and the ergosterol peak cannot otherwise be resolved. Using a 2-h acetylation at 601C, recoveries of 10 and 25 ppm added ergosterol were 96.3% and 94.0%, respectively (n ¼ 4; relative SD=3.8% in each case). Results using a convenient overnight acetylation at 371C were similar and not significantly different. The limit of detection was approximately 0.3 ppm. Crown Copyright r 2002 Published by Elsevier Science Ltd. All rights reserved. Keywords: Rapeseed; Canola; Ergosterol; Chromatography; Spoilage
1. Introduction Canola (Brassica napus L.) is a type of rapeseed producing oil with low erucic acid levels, and meal with a low content of glucosinolates. Canola has become an important oilseed crop in western Canada and Australia, while traditional types of rapeseed continue to be grown in other countries such as China, India, Germany, France, and Britain. Although 7.1 million tonnes of canola were produced in Canada during 1999, 1.4 million tonnes were stored in inland and coastal *Corresponding author. Tel.: +1-204-983-1469; fax: +1-204-983-4604. E-mail address: [email protected] (D. Abramson). 0022-474X/02/$ - see front matter Crown Copyright r 2002 Published by Elsevier Science Ltd. All rights reserved. PII: S 0 0 2 2 - 4 7 4 X ( 0 1 ) 0 0 0 5 2 - 2
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D. Abramson, D.M. Smith / Journal of Stored Products Research 39 (2003) 185–191
elevators, or on farms, for a year or more. Even at a relatively low moisture content such as 10%, canola is susceptible to spoilage by fungi if temperatures are high enough and the storage period is long enough (Mills and Sinha, 1980). Several methods have been proposed to assess fungal contamination of canola and other crops. The determination of fungal biomass is always a problem when the physical separation of fungus from substrate is impossible, and one solution is to determine fungus-specific chemical constituents and their relation to mycelial dry mass. Both glucosamine and ergosterol have been examined in this regard, but ergosterol is considered more accurate (Seitz et al., 1979; Schwadorf and Muller, . 1989; Johnson and McGill, 1990). Using saponification, both esterified and free ergosterol can be estimated, and this so-called ‘‘total’’ ergosterol measurement can serve as a more sensitive estimation of fungal biomass than free ergosterol alone (Seitz et al., 1979; Tothill et al., 1992). Both liquid chromatography (LC) and gas chromatography (GC) have been used to detect total ergosterol in fungal cultures and plant material (Piironen et al., 2000). Seitz et al. (1979) used reverse-phase LC with absorbency detection at 282 nm to assay fungal cultures, but method performance parameters such as recovery of added ergosterol were not reported. Reverse-phase LC was also used in studies on decomposing plant material by Newell et al. (1988) who found 95–107% recoveries of ergosterol, and by Gessner and Schmitt (1996) who reported 85–98%. In studies on various animal feed constituents, Schwadorf and Muller . (1989) reported 97–102% recoveries using normal-phase (silica gel) LC. In other crops used as oil sources, Kaminski et al. (1985) used GC to assay ergosterol in high-oil corn, and Dhingra et al. (1998) determined ergosterol in soybeans using a reverse-phase LC procedure, but method performance was not evaluated in either case. Through incorporating and modifying various steps from the above methods, a novel procedure resulted for determining total ergosterol in canola by reverse-phase LC. The strong ultraviolet absorbency of ergosterol at 282 nm makes its assay possible without interference from other phytosterols, providing that care is taken to separate the analyte from other substances in the matrix. The very high oil content of canola, typically 44–46% (Daun et al., 1993), precluded direct saponification. A contaminant co-chromatographing with free ergosterol required that the purified extracts be acetylated, and that ergosterol be quantitated as the acetate.
2. Materials and methods Seed-grade canola (cultivar Lima Grain 3295) was ground for 30 s in a rotary mill (Model M-2, F. Stein Labs Inc., Atchison, KS, USA) and 4.0 g were extracted by refluxing for 1 h in 50 ml methanol. After cooling, the mixture was filtered through Whatman 802 paper, and the filtrate volume adjusted to 60 ml with methanol. For saponification, 10 ml methanol containing 2 g dissolved potassium hydroxide was added, and the mixture refluxed for 30 min. After cooling, filtration through Whatman 802 paper, rinsing three times with 5 ml methanol, and addition of 15 ml water to the filtrate, the mixture was extracted twice with 50 ml n-hexane. The n-hexane extracts were combined, dried over 15 g anhydrous sodium sulfate, and 12.5 ml removed and taken to dryness under a nitrogen stream at 401C.
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The residue was reconstituted to 2 ml in cyclohexane, added to a 3-ml solid-phase extraction (SPE) cartridge containing 500 mg silica gel (Sep Pack WAT 020810, Waters Inc., Millford, MA, USA), and rinsed onto the cartridge with two further portions of 2 ml cyclohexane. The cartridge was washed with 6 ml carbon tetrachloride and aspirated to dryness (15 min). Both the cyclohexane and carbon tetrachloride were dried with anhydrous sodium sulfate before use. The silica cartridge was then eluted with 2.5 ml acetone, the eluate taken to dryness under nitrogen at 301C, and then reconstituted in 0.5 ml pyridine. For acetylation, 0.5 ml acetic anhydride was added, and after 2 h at 601C under dark conditions, the mixture was taken to dryness under nitrogen at 451C. For LC analysis, the purified residue was taken up in 1 ml acetonitrile with sonication, and clarified by centrifugation at 15,600 g for 3 min. Aliquots of 50 mL were injected into an LC system with a C-18 reverse-phase column (Lichrospher Si-100 RP-18, 25 cm long, 4 mm diameter, 10 mm particles, E. Merck, Darmstadt, Germany) at 451C, and a diode-array spectrophotometric detector set at 282 nm. Using methanol at 1.0 ml/min as the mobile phase, ergosterol acetate was consistently eluted at 13.6 min. For estimation of ergosterol in the canola residues, standards of ergosterol (>98%, Fluka Chemie AG, Buchs, Switzerland) were made up in absolute ethanol, quantitated by absorbance at 282 nm using e=12,150 (Shaw and Jeffrey, 1953), and dried and acetylated as above. Prior to recovery studies, it was necessary to determine background levels of total ergosterol in non-moldy canola following the above procedure. For recovery studies on added ergosterol, known amounts were added to the ground canola by adding appropriate volumes of ergosterol standard solutions (100 mg/ml in dichloromethane), and allowing the solvent to evaporate. Within 1 h of ergosterol addition, this spiked sample was extracted and treated following the described procedure. Background levels were subtracted in all recovery determinations. Background levels and recoveries were also studied using an alternative overnight acetylation procedure, in which samples were reacted at 371C for 17 h under dark conditions.
3. Results and discussion The acetylation procedure afforded liquid chromatograms with baseline separation of ergosterol acetate from the other components of the mixture at background levels (Fig. 1), with 10 ppm ergosterol added (Fig. 2), and with 25 ppm ergosterol added. Ergosterol acetate eluted at 13.6070.03 min. In the background-level samples, the diode-array absorbance spectrum between 240 and 310 nm for the well-separated peak at 13.60 min was identical to that of standard ergosterol acetate. The limit of detection would be approximately 0.3 ppm if canola with such abnormally low background levels could be found. The actual background levels in canola were found to be 2.8–3.1 ppm (Table 1). The 10 and 25 ppm spiking levels reflect ergosterol levels expected in moldy canola based on data from other high-lipid matrices. Between 9 and 58 ppm of ergosterol has been reported in moldy corn (Kaminski et al., 1985), while Dhingra et al. (1998) found 6–7 ppm in moldy soybeans. As mentioned in the introduction, acetylation was necessary to separate ergosterol from an unidentified contaminant which co-eluted during LC analysis. A chromatogram of non-acetylated canola extract containing 10 ppm added ergosterol is shown in Fig. 3. Under the chromatography
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Fig. 1. Determination of background level of ergosterol in canola after acetylation. Aliquots of 50 mL were injected onto a 25 cm long column of Merck Lichrospher RP-18, 10 mm particle diameter, and eluted with methanol 1.0 ml/min at 451C. Absorbance was monitored at 282 nm.
conditions described above, ergosterol eluted at 8.4770.05 min. Some separation of ergosterol from the major contaminant eluting at 8.79 min is evident, but resolution of the two peaks is not sufficient for consistent quantitation. At background levels of approximately 3 ppm (Table 1), ergosterol could not be resolved from the contaminant and appears as a slight shoulder on the peak. If this entire peak were erroneously quantitated as ergosterol, then background levels of ergosterol in non-moldy canola would appear to be 35–40 ppm. Diode-array absorbance spectra between 240 and 310 nm for standard ergosterol, and for the major contaminant, were dissimilar, with the characteristic ergosterol double peak at 272 and 282 nm missing in the contaminant peak. The mean background levels of ergosterol in canola, and recoveries for 10 and 25 ppm added ergosterol, are indicated in Table 1. A 2-h saponification at 601C and a 17-h overnight saponification at 371C were both employed, and gave measurable recoveries. Background levels were calculated separately for each procedure. Using Student’s t-test, the two saponification procedures produced no significant differences (P > 0:05) between mean background levels of ergosterol, between mean recoveries of 10 ppm added ergosterol, or between mean recoveries of 25 ppm added ergosterol. Although lengthy, our method produced high recoveries of ergosterol from the canola matrix, with a high level of precision among replicate assays. Comparison of method performance with other ergosterol assays in other high oil content crops is difficult, since no specific recovery data have been cited for these methods (Kaminski et al., 1985; Dhingra et al., 1998). In their assay for ergosterol in 24 feedstuff ingredients, which included some cereal and
D. Abramson, D.M. Smith / Journal of Stored Products Research 39 (2003) 185–191
189
Fig. 2. Determination of ergosterol as the acetate in canola containing 10 ppm added ergosterol. Aliquots of 50 mL were injected onto a 25 cm long column of Merck Lichrospher RP-18, 10 mm particle diameter, and eluted with methanol 1.0 ml/min at 451C. Absorbance was monitored at 282 nm. Table 1 Recoveries of added ergosterol from canola, using two different acetylation conditions Acetylation conditions 371C, 371C, 371C, 601C, 601C, 601C,
17 h 17 h 17 h 2h 2h 2h
Mean ergosterol Added (ppm)
Found (ppm)
0 10 25 0 10 25
2.8b 12.3 25.8 3.1b 12.7 26.6
Percentage recovery of added ergosterola
Percentage relative SD
n
F 94.8 92.0 F 96.3 94.0
7.0 10.6 6.4 8.1 3.8 3.8
4 4 4 4 4 4
level found background 100: level added b Background levels. a
% recovery ¼
oilseed matrices, Schwadorf and Muller . (1989) reported recoveries between 96.7% and 102.2%, but no specific recoveries for oilseeds, or measure of variation among replicates for oilseeds, was given. Prior to saponification, a methanol extraction was necessary to leave behind most of the canola oil. Early attempts involving direct saponification of the ground canola with 3–8% potassium
190
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Fig. 3. Chromatogram of a non-acetylated canola extract containing 10 ppm added ergosterol. Aliquots of 50 mL were injected onto a 25 cm long column of Merck Lichrospher RP-18, 10 mm particle diameter, and eluted with methanol 1.0 ml/min at 451C. Absorbance was monitored at 282 nm.
hydroxide in methanol yielded a product which contained large amounts of fatty acids. These interfered with all subsequent efforts at purification, and recoveries o45% of added ergosterol were observed. The silica gel SPE technique proved to be a rapid and convenient way to separate polar analytes from non-polar lipid contaminants (Bello, 1992). Because water inactivates this adsorbent, care should be taken to dry the n-hexane extracts with sodium sulfate, and to completely evaporate the solvent to produce a dry-film residue in the collection vials. Early attempts at silica gel SPE using the n-hexane extract of the saponification mixture directly, without the drying, evaporation and reconstitution steps, gave ergosterol recoveries of only approximately 65–70%. This was due to water entrainment from the extraction step, inactivation of the silica, and partial breakthrough of the analyte. It might be possible to develop a shorter assay for ergosterol in canola eliminating the SPE step and acetylation, but this procedure would require normal-phase silica LC for quantitation. Any potential advantage of developing a method based on normal-phase silica LC should be considered against three distinct disadvantages. First, small amounts of moisture in normal-phase LC running solvents such as hexane can significantly alter retention times. Hexane can contain up to 0.2% water, absorbs moisture from the air under humid ambient conditions (Dolan, 1998), and would require special drying treatment. Second, exhaustive regeneration procedures (Schwadorf and Muller, . 1989) are necessary to restore silica column performance and accurate retention
D. Abramson, D.M. Smith / Journal of Stored Products Research 39 (2003) 185–191
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times, which are not necessary on reverse-phase columns. Third, the use of relatively volatile normal-phase LC solvents such as dichloromethane and hexane presents hazards to laboratory workers from both flammability and inhalation toxicity, and additional fume hood space would be needed for LC equipment and LC solvent preparation. The proposed method provides a precise and quantitative procedure for assessing ergosterol content and fungal contamination in agricultural products having a high lipid content, such as canola. Other high-oil crops could be assayed with the proposed method, particularly flaxseed, with an oil content of 41–44%, and sunflower seed, with an oil content of 42–48% (Daun et al., 1993). A preliminary analysis without acetylation would be necessary to indicate if any interfering peaks co-chromatographing with ergosterol were present, as in canola. If interfering material was not present, the procedure could be shortened by elimination of the acetylation step. Acknowledgements The authors thank Carl Pronyk, Biosystems Engineering Department, University of Manitoba, for providing ground samples of canola. This report is Contribution Number 1820 of the Cereal Research Centre. References Bello, A.C., 1992. Rapid isolation of the sterol fraction in edible oils using a silica cartridge. Journal of the AOAC International 75, 1120–1123. Daun, J.K., Buhr, N., Diosady, L.L., Mills, J.T., Mag, T., 1993. OilseedsFprocessing. In: Grains and Oilseeds: Handling Marketing Processing, Vol. 2. Canadian International Grains Institute, Winnipeg, pp. 883–935. Dhingra, O.D., Jham, G., Napoleao, I.T., 1998. Ergosterol accumulation and oil quality changes in stored soybean invaded by Aspergillus ruber (A. glaucus group). Mycopathologia 143, 85–91. Dolan, J.W., 1998. Column packingFwhat is at the bottom of it? LC–GC 16, 350–354. Gessner, M.O., Schmitt, A.L., 1996. Use of solid-phase extraction to determine ergosterol concentrations in plant tissue colonized by fungi. Applied and Environmental Microbiology 62, 415–419. Johnson, B.N., McGill, W.B., 1990. Comparison of ergosterol and chitin as quantitative estimates of mycorrhizal infection and Pinus contorta response to inoculation. Canadian Journal of Forestry Research 20, 1125–1131. Kaminski, E., Wasowicz, E., Gruchala, L., 1985. Determination of ergosterol by GLC in fat-containing raw material as a criterion of mold invasion. Developments in Food Science 11, 319–326. Mills, J.T., Sinha, R.N., 1980. Safe storage periods for farm-stored rapeseed based on mycological and biochemical assessment. Phytopathology 70, 541–547. Newell, S.Y., Arsuffi, T.L., Fallon, R.D., 1988. Fundamental procedures for determining ergosterol content of decaying plant material by liquid chromatography. Applied and Environmental Microbiology 54, 1876–1879. Piironen, V., Lindsay, D.G., Meittinen, T.A., Toivo, J., Lampi, A.-M., 2000. Plant sterols: biosynthesis, biological function and their importance to human nutrition. Journal of the Science of Food and Agriculture 80, 939–966. Schwadorf, K., Muller, . H.-M., 1989. Determination of ergosterol in cereals, mixed feed components, and mixed feeds by liquid chromatography. Journal of the AOAC International 72, 457–462. Seitz, L.M., Sauer, D.B., Mohr, H.E., Hubbard, J.D., 1979. Ergosterol as a measure of fungal growth. Phytopathology 69, 1202–1203. Shaw, W.H.C., Jeffrey, J.P., 1953. The determination of ergosterol in yeast. Analyst 78, 509–514. Tothill, I.E., Harris, D., Magan, N., 1992. The relationship between fungal growth and ergosterol content in wheat grain. Mycological Research 96, 965–970.
Determination of ergosterol in canola (Brassica napus L.) by liquid chromatography D. Abramson*, D.M. Smith Cereal Research Centre, Agriculture and Agri-Food Canada, 195 Dafoe Road, Winnipeg, Man., Canada R3T 2M9 Accepted 22 October 2001
Abstract A sensitive and precise method is described to assay the oilseed canola (Brassica napus L.) for ergosterol, a fungal metabolite indicating spoilage. The ground seed is refluxed in methanol, and the methanol extract is saponified with potassium hydroxide. After addition of water, the mixture is partitioned into n-hexane. The n-hexane extract is dried, reconstituted, and applied to a silica solid-phase extraction cartridge, which is then washed with carbon tetrachloride, and eluted with acetone. The acetone eluate is acetylated, and the ergosterol determined as the acetate by liquid chromatography using a reverse-phase column and absorbency detection at 282 nm. Acetylation is necessary since an unidentified constituent of the matrix cochromatographs with free ergosterol, and the ergosterol peak cannot otherwise be resolved. Using a 2-h acetylation at 601C, recoveries of 10 and 25 ppm added ergosterol were 96.3% and 94.0%, respectively (n ¼ 4; relative SD=3.8% in each case). Results using a convenient overnight acetylation at 371C were similar and not significantly different. The limit of detection was approximately 0.3 ppm. Crown Copyright r 2002 Published by Elsevier Science Ltd. All rights reserved. Keywords: Rapeseed; Canola; Ergosterol; Chromatography; Spoilage
1. Introduction Canola (Brassica napus L.) is a type of rapeseed producing oil with low erucic acid levels, and meal with a low content of glucosinolates. Canola has become an important oilseed crop in western Canada and Australia, while traditional types of rapeseed continue to be grown in other countries such as China, India, Germany, France, and Britain. Although 7.1 million tonnes of canola were produced in Canada during 1999, 1.4 million tonnes were stored in inland and coastal *Corresponding author. Tel.: +1-204-983-1469; fax: +1-204-983-4604. E-mail address: [email protected] (D. Abramson). 0022-474X/02/$ - see front matter Crown Copyright r 2002 Published by Elsevier Science Ltd. All rights reserved. PII: S 0 0 2 2 - 4 7 4 X ( 0 1 ) 0 0 0 5 2 - 2
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elevators, or on farms, for a year or more. Even at a relatively low moisture content such as 10%, canola is susceptible to spoilage by fungi if temperatures are high enough and the storage period is long enough (Mills and Sinha, 1980). Several methods have been proposed to assess fungal contamination of canola and other crops. The determination of fungal biomass is always a problem when the physical separation of fungus from substrate is impossible, and one solution is to determine fungus-specific chemical constituents and their relation to mycelial dry mass. Both glucosamine and ergosterol have been examined in this regard, but ergosterol is considered more accurate (Seitz et al., 1979; Schwadorf and Muller, . 1989; Johnson and McGill, 1990). Using saponification, both esterified and free ergosterol can be estimated, and this so-called ‘‘total’’ ergosterol measurement can serve as a more sensitive estimation of fungal biomass than free ergosterol alone (Seitz et al., 1979; Tothill et al., 1992). Both liquid chromatography (LC) and gas chromatography (GC) have been used to detect total ergosterol in fungal cultures and plant material (Piironen et al., 2000). Seitz et al. (1979) used reverse-phase LC with absorbency detection at 282 nm to assay fungal cultures, but method performance parameters such as recovery of added ergosterol were not reported. Reverse-phase LC was also used in studies on decomposing plant material by Newell et al. (1988) who found 95–107% recoveries of ergosterol, and by Gessner and Schmitt (1996) who reported 85–98%. In studies on various animal feed constituents, Schwadorf and Muller . (1989) reported 97–102% recoveries using normal-phase (silica gel) LC. In other crops used as oil sources, Kaminski et al. (1985) used GC to assay ergosterol in high-oil corn, and Dhingra et al. (1998) determined ergosterol in soybeans using a reverse-phase LC procedure, but method performance was not evaluated in either case. Through incorporating and modifying various steps from the above methods, a novel procedure resulted for determining total ergosterol in canola by reverse-phase LC. The strong ultraviolet absorbency of ergosterol at 282 nm makes its assay possible without interference from other phytosterols, providing that care is taken to separate the analyte from other substances in the matrix. The very high oil content of canola, typically 44–46% (Daun et al., 1993), precluded direct saponification. A contaminant co-chromatographing with free ergosterol required that the purified extracts be acetylated, and that ergosterol be quantitated as the acetate.
2. Materials and methods Seed-grade canola (cultivar Lima Grain 3295) was ground for 30 s in a rotary mill (Model M-2, F. Stein Labs Inc., Atchison, KS, USA) and 4.0 g were extracted by refluxing for 1 h in 50 ml methanol. After cooling, the mixture was filtered through Whatman 802 paper, and the filtrate volume adjusted to 60 ml with methanol. For saponification, 10 ml methanol containing 2 g dissolved potassium hydroxide was added, and the mixture refluxed for 30 min. After cooling, filtration through Whatman 802 paper, rinsing three times with 5 ml methanol, and addition of 15 ml water to the filtrate, the mixture was extracted twice with 50 ml n-hexane. The n-hexane extracts were combined, dried over 15 g anhydrous sodium sulfate, and 12.5 ml removed and taken to dryness under a nitrogen stream at 401C.
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The residue was reconstituted to 2 ml in cyclohexane, added to a 3-ml solid-phase extraction (SPE) cartridge containing 500 mg silica gel (Sep Pack WAT 020810, Waters Inc., Millford, MA, USA), and rinsed onto the cartridge with two further portions of 2 ml cyclohexane. The cartridge was washed with 6 ml carbon tetrachloride and aspirated to dryness (15 min). Both the cyclohexane and carbon tetrachloride were dried with anhydrous sodium sulfate before use. The silica cartridge was then eluted with 2.5 ml acetone, the eluate taken to dryness under nitrogen at 301C, and then reconstituted in 0.5 ml pyridine. For acetylation, 0.5 ml acetic anhydride was added, and after 2 h at 601C under dark conditions, the mixture was taken to dryness under nitrogen at 451C. For LC analysis, the purified residue was taken up in 1 ml acetonitrile with sonication, and clarified by centrifugation at 15,600 g for 3 min. Aliquots of 50 mL were injected into an LC system with a C-18 reverse-phase column (Lichrospher Si-100 RP-18, 25 cm long, 4 mm diameter, 10 mm particles, E. Merck, Darmstadt, Germany) at 451C, and a diode-array spectrophotometric detector set at 282 nm. Using methanol at 1.0 ml/min as the mobile phase, ergosterol acetate was consistently eluted at 13.6 min. For estimation of ergosterol in the canola residues, standards of ergosterol (>98%, Fluka Chemie AG, Buchs, Switzerland) were made up in absolute ethanol, quantitated by absorbance at 282 nm using e=12,150 (Shaw and Jeffrey, 1953), and dried and acetylated as above. Prior to recovery studies, it was necessary to determine background levels of total ergosterol in non-moldy canola following the above procedure. For recovery studies on added ergosterol, known amounts were added to the ground canola by adding appropriate volumes of ergosterol standard solutions (100 mg/ml in dichloromethane), and allowing the solvent to evaporate. Within 1 h of ergosterol addition, this spiked sample was extracted and treated following the described procedure. Background levels were subtracted in all recovery determinations. Background levels and recoveries were also studied using an alternative overnight acetylation procedure, in which samples were reacted at 371C for 17 h under dark conditions.
3. Results and discussion The acetylation procedure afforded liquid chromatograms with baseline separation of ergosterol acetate from the other components of the mixture at background levels (Fig. 1), with 10 ppm ergosterol added (Fig. 2), and with 25 ppm ergosterol added. Ergosterol acetate eluted at 13.6070.03 min. In the background-level samples, the diode-array absorbance spectrum between 240 and 310 nm for the well-separated peak at 13.60 min was identical to that of standard ergosterol acetate. The limit of detection would be approximately 0.3 ppm if canola with such abnormally low background levels could be found. The actual background levels in canola were found to be 2.8–3.1 ppm (Table 1). The 10 and 25 ppm spiking levels reflect ergosterol levels expected in moldy canola based on data from other high-lipid matrices. Between 9 and 58 ppm of ergosterol has been reported in moldy corn (Kaminski et al., 1985), while Dhingra et al. (1998) found 6–7 ppm in moldy soybeans. As mentioned in the introduction, acetylation was necessary to separate ergosterol from an unidentified contaminant which co-eluted during LC analysis. A chromatogram of non-acetylated canola extract containing 10 ppm added ergosterol is shown in Fig. 3. Under the chromatography
188
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Fig. 1. Determination of background level of ergosterol in canola after acetylation. Aliquots of 50 mL were injected onto a 25 cm long column of Merck Lichrospher RP-18, 10 mm particle diameter, and eluted with methanol 1.0 ml/min at 451C. Absorbance was monitored at 282 nm.
conditions described above, ergosterol eluted at 8.4770.05 min. Some separation of ergosterol from the major contaminant eluting at 8.79 min is evident, but resolution of the two peaks is not sufficient for consistent quantitation. At background levels of approximately 3 ppm (Table 1), ergosterol could not be resolved from the contaminant and appears as a slight shoulder on the peak. If this entire peak were erroneously quantitated as ergosterol, then background levels of ergosterol in non-moldy canola would appear to be 35–40 ppm. Diode-array absorbance spectra between 240 and 310 nm for standard ergosterol, and for the major contaminant, were dissimilar, with the characteristic ergosterol double peak at 272 and 282 nm missing in the contaminant peak. The mean background levels of ergosterol in canola, and recoveries for 10 and 25 ppm added ergosterol, are indicated in Table 1. A 2-h saponification at 601C and a 17-h overnight saponification at 371C were both employed, and gave measurable recoveries. Background levels were calculated separately for each procedure. Using Student’s t-test, the two saponification procedures produced no significant differences (P > 0:05) between mean background levels of ergosterol, between mean recoveries of 10 ppm added ergosterol, or between mean recoveries of 25 ppm added ergosterol. Although lengthy, our method produced high recoveries of ergosterol from the canola matrix, with a high level of precision among replicate assays. Comparison of method performance with other ergosterol assays in other high oil content crops is difficult, since no specific recovery data have been cited for these methods (Kaminski et al., 1985; Dhingra et al., 1998). In their assay for ergosterol in 24 feedstuff ingredients, which included some cereal and
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189
Fig. 2. Determination of ergosterol as the acetate in canola containing 10 ppm added ergosterol. Aliquots of 50 mL were injected onto a 25 cm long column of Merck Lichrospher RP-18, 10 mm particle diameter, and eluted with methanol 1.0 ml/min at 451C. Absorbance was monitored at 282 nm. Table 1 Recoveries of added ergosterol from canola, using two different acetylation conditions Acetylation conditions 371C, 371C, 371C, 601C, 601C, 601C,
17 h 17 h 17 h 2h 2h 2h
Mean ergosterol Added (ppm)
Found (ppm)
0 10 25 0 10 25
2.8b 12.3 25.8 3.1b 12.7 26.6
Percentage recovery of added ergosterola
Percentage relative SD
n
F 94.8 92.0 F 96.3 94.0
7.0 10.6 6.4 8.1 3.8 3.8
4 4 4 4 4 4
level found background 100: level added b Background levels. a
% recovery ¼
oilseed matrices, Schwadorf and Muller . (1989) reported recoveries between 96.7% and 102.2%, but no specific recoveries for oilseeds, or measure of variation among replicates for oilseeds, was given. Prior to saponification, a methanol extraction was necessary to leave behind most of the canola oil. Early attempts involving direct saponification of the ground canola with 3–8% potassium
190
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Fig. 3. Chromatogram of a non-acetylated canola extract containing 10 ppm added ergosterol. Aliquots of 50 mL were injected onto a 25 cm long column of Merck Lichrospher RP-18, 10 mm particle diameter, and eluted with methanol 1.0 ml/min at 451C. Absorbance was monitored at 282 nm.
hydroxide in methanol yielded a product which contained large amounts of fatty acids. These interfered with all subsequent efforts at purification, and recoveries o45% of added ergosterol were observed. The silica gel SPE technique proved to be a rapid and convenient way to separate polar analytes from non-polar lipid contaminants (Bello, 1992). Because water inactivates this adsorbent, care should be taken to dry the n-hexane extracts with sodium sulfate, and to completely evaporate the solvent to produce a dry-film residue in the collection vials. Early attempts at silica gel SPE using the n-hexane extract of the saponification mixture directly, without the drying, evaporation and reconstitution steps, gave ergosterol recoveries of only approximately 65–70%. This was due to water entrainment from the extraction step, inactivation of the silica, and partial breakthrough of the analyte. It might be possible to develop a shorter assay for ergosterol in canola eliminating the SPE step and acetylation, but this procedure would require normal-phase silica LC for quantitation. Any potential advantage of developing a method based on normal-phase silica LC should be considered against three distinct disadvantages. First, small amounts of moisture in normal-phase LC running solvents such as hexane can significantly alter retention times. Hexane can contain up to 0.2% water, absorbs moisture from the air under humid ambient conditions (Dolan, 1998), and would require special drying treatment. Second, exhaustive regeneration procedures (Schwadorf and Muller, . 1989) are necessary to restore silica column performance and accurate retention
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times, which are not necessary on reverse-phase columns. Third, the use of relatively volatile normal-phase LC solvents such as dichloromethane and hexane presents hazards to laboratory workers from both flammability and inhalation toxicity, and additional fume hood space would be needed for LC equipment and LC solvent preparation. The proposed method provides a precise and quantitative procedure for assessing ergosterol content and fungal contamination in agricultural products having a high lipid content, such as canola. Other high-oil crops could be assayed with the proposed method, particularly flaxseed, with an oil content of 41–44%, and sunflower seed, with an oil content of 42–48% (Daun et al., 1993). A preliminary analysis without acetylation would be necessary to indicate if any interfering peaks co-chromatographing with ergosterol were present, as in canola. If interfering material was not present, the procedure could be shortened by elimination of the acetylation step. Acknowledgements The authors thank Carl Pronyk, Biosystems Engineering Department, University of Manitoba, for providing ground samples of canola. This report is Contribution Number 1820 of the Cereal Research Centre. References Bello, A.C., 1992. Rapid isolation of the sterol fraction in edible oils using a silica cartridge. Journal of the AOAC International 75, 1120–1123. Daun, J.K., Buhr, N., Diosady, L.L., Mills, J.T., Mag, T., 1993. OilseedsFprocessing. In: Grains and Oilseeds: Handling Marketing Processing, Vol. 2. Canadian International Grains Institute, Winnipeg, pp. 883–935. Dhingra, O.D., Jham, G., Napoleao, I.T., 1998. Ergosterol accumulation and oil quality changes in stored soybean invaded by Aspergillus ruber (A. glaucus group). Mycopathologia 143, 85–91. Dolan, J.W., 1998. Column packingFwhat is at the bottom of it? LC–GC 16, 350–354. Gessner, M.O., Schmitt, A.L., 1996. Use of solid-phase extraction to determine ergosterol concentrations in plant tissue colonized by fungi. Applied and Environmental Microbiology 62, 415–419. Johnson, B.N., McGill, W.B., 1990. Comparison of ergosterol and chitin as quantitative estimates of mycorrhizal infection and Pinus contorta response to inoculation. Canadian Journal of Forestry Research 20, 1125–1131. Kaminski, E., Wasowicz, E., Gruchala, L., 1985. Determination of ergosterol by GLC in fat-containing raw material as a criterion of mold invasion. Developments in Food Science 11, 319–326. Mills, J.T., Sinha, R.N., 1980. Safe storage periods for farm-stored rapeseed based on mycological and biochemical assessment. Phytopathology 70, 541–547. Newell, S.Y., Arsuffi, T.L., Fallon, R.D., 1988. Fundamental procedures for determining ergosterol content of decaying plant material by liquid chromatography. Applied and Environmental Microbiology 54, 1876–1879. Piironen, V., Lindsay, D.G., Meittinen, T.A., Toivo, J., Lampi, A.-M., 2000. Plant sterols: biosynthesis, biological function and their importance to human nutrition. Journal of the Science of Food and Agriculture 80, 939–966. Schwadorf, K., Muller, . H.-M., 1989. Determination of ergosterol in cereals, mixed feed components, and mixed feeds by liquid chromatography. Journal of the AOAC International 72, 457–462. Seitz, L.M., Sauer, D.B., Mohr, H.E., Hubbard, J.D., 1979. Ergosterol as a measure of fungal growth. Phytopathology 69, 1202–1203. Shaw, W.H.C., Jeffrey, J.P., 1953. The determination of ergosterol in yeast. Analyst 78, 509–514. Tothill, I.E., Harris, D., Magan, N., 1992. The relationship between fungal growth and ergosterol content in wheat grain. Mycological Research 96, 965–970.