GC and On-line LC-GC - American Chemical Society

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Chapter 13

GC and On-line LC-GC: Useful Tools for the Qualitative and Quantitative Analysis of Phytosterols and Their Esters Rebecca Esche, Birgit Scholz, and Karl-Heinz Engel* Technische Universität München, Lehrstuhl für Allgemeine Lebensmitteltechnologie, Maximus-von-Imhof-Forum 2, D-85350 Freising-Weihenstephan, Germany *E-Mail: [email protected].

Phytosterols/-stanols and their esters possess several health-benefits such as cholesterol-lowering and anti-oxidative properties. For the qualitative and quantitative analysis of individual free sterols/stanols and intact steryl/stanyl esters in various food matrices different approaches were established. A combination of GC-based analysis and fractionation via solid-phase extraction was applied to investigate the natural variability of these compounds in cereal grains. On-line LC-GC was demonstrated to be a useful tool for the rapid analysis of stanyl fatty acid esters in enriched dairy foods as well as for the simultaneous analysis of free sterols/stanols, steryl/stanyl fatty acid esters, and other minor lipids in edible plant oils and nuts.

Introduction Phytosterols/-stanols are bioactive secondary plant metabolites occurring in free form, esterified with fatty acids or phenolic acids (Figure 1), and as glycosides or acylated glycosides (1). The nutritional interest in these compounds mainly arises from their cholesterol-lowering properties (2, 3). A total intake of 2 g plant sterols/stanols per day can reduce the levels of low-density lipoprotein cholesterol in hypercholesterolemic patients by up to 10 % (4). For this purpose, a broad spectrum of foods such as spread, margarine, yogurt, or milk © 2014 American Chemical Society In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.


is currently enriched with mixtures of plant steryl/stanyl fatty acid esters. In the European Union, these mixtures are specified concerning the distributions of esterified sterols/stanols and fatty acids (5–7). Vegetable oils can be used as sources to provide phytosterols/-stanols and the fatty acid mixtures used for the esterification; tall oil is also a possible source to obtain sterols/-stanols. The following profile of free or esterified sterols/stanols is considered as acceptable in general for the incorporation into enriched food products in the European Union: up to 80% β-sitosterol, 40% campesterol, 30% stigmasterol, 15% sitostanol, 5% campestanol, 3% brassicasterol and 3% other phytosterols (6). Among natural foods, cereals, nuts, and edible plant oils are particularly rich sources of free sterols/stanols and steryl/stanyl esters (1).

Figure 1. Representative structures of a free sterol and steryl esters: (A) sitosterol, (B) trans-sitosteryl ferulate, and (C) sitosteryl oleate.

Previously, qualitative and quantitative investigations have been mainly based on the analysis of total sterols/stanols determined after alkaline hydrolysis of the lipids. As a result, information on the contents and compositions particularly of individual intact steryl/stanyl fatty acid esters is rare. Therefore, analytical approaches are needed allowing both the authentication of foods enriched with steryl/stanyl fatty acid esters as well as the determination of naturally occurring free sterols/stanols and their intact esters. Recently, high temperature capillary gas chromatography was demonstrated to be suitable for the analysis of complex mixtures of steryl/stanyl fatty acid esters (8, 9). On this basis, three methodologies were established enabling the analysis of stanyl fatty acid esters in enriched dairy foods as well as the comprehensive analysis of free sterols/stanols and individual steryl/stanyl fatty acid and phenolic acid esters in cereals, edible plant oils, and nuts. 258 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.

Experimental Section


Materials and Chemicals The cereal grains, edible vegetable oils, and nuts were obtained in local stores (Freising, Germany). The cheese-based spread enriched with stanyl fatty acid esters (“Benecol naturalny-krem kanapkowy” produced by Raisio Sp. Z o.o/Bahca Polska Sp z o.o, Warzawa, Poland) was purchased in a supermarket in Poland. A mixture of plant stanyl fatty acid esters (“plant stanol ester, STAEST 115”) was provided by Raisio Group (Raisio, Finland). Reference compounds of steryl/stanyl fatty acid esters and steryl/stanyl phenolic acid esters were synthesized according to previously described procedures (8, 10). All other chemicals and solvents were obtained from Sigma Aldrich (Steinheim, Germany), VWR International (Darmstadt, Germany), Evonik Industries AG (Essen, Germany) or Acros Organics (Morris Plains, NJ, U.S.A.). Sample Preparation Enriched Dairy Foods After homogenization and addition of the internal standard (IS, cholesteryl16:0), the samples were subjected to an acid digestion step with hydrochloric acid (25 %) at 130 °C for 45 min. The lipids were extracted with a mixture of n-hexane/ methyl tert-butyl ether (MTBE; 3:2, v/v), and the extracts were directly used for on-line LC-GC analysis (11).

Cereal Grains Lipids were extracted from ground and freeze-dried cereal grains after the addition of the internal standards (5α-cholestan-3β-ol, cholesteryl-16:0, and cholestanyl p-coumarate) with of a mixture of n-hexane/dichloromethane (1:1, v/v) under stirring for 1 h at room temperature. The solvent was removed by rotary evaporation and 100 mg of the obtained oil was dissolved in 10 mL of n-hexane; 1 mL of that solution was used for SPE (10).

Edible Plant Oils and Nuts The commercially obtained edible plant oils were spiked with the internal standards C17:0, 5α-cholestan-3β-ol, cholesteryl-16:0, and trans-cholestanyl ferulate. The nut samples were ground to a fine powder and lipids were extracted after the addition of the internal standards (5α-cholestan-3β-ol and cholesteryl-16:0) with of a mixture of n-hexane/dichloromethane (1:1, v/v) under stirring for 1 h at room temperature. The solvent was removed by rotary evaporation (12). 259 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.


Fifty milligrams of the edible plant oils and of the obtained nut oils, respectively, were dissolved in 5 mL of n-hexane. An aliquot (250 µL) of the solution was dried by a gentle stream of nitrogen, and the residue was silylated with 75 µL of pyridine and 150 µL of N,O-bis(trimethylsilyl)trifluoroacetamide/ trimethyl-chlorosilane (99:1, v/v) at 80 °C for 20 min. After removal of the reagents, the residue was dissolved in 250 µL of n-hexane/MTBE/2-propanol (96:4:0.1, v/v/v) and used for on-line LC-GC analysis (12). Gas Chromatography-Flame Ionization Detection (GC-FID) and Gas Chromatography-Mass Spectrometry (GC-MS) Separations were performed using a gas chromatograph equipped with an FID (Agilent Technologies Instrument 6890N, Böblingen, Germany). The sample solutions (1 μL) were injected onto a 30 m × 0.25 mm i.d., 0.1 µm film, trifluoropropylmethyl polysiloxane capillary column (Rtx-200MS, Restek, Bad Homburg, Germany). The temperature of the injector was set to 280 °C. Hydrogen was used as carrier gas with constant flow (1.5 mL/min) and the split flow was set to 11.2 mL/min. The oven temperature program was as follows: initial temperature 100 °C, 15 °C/min to 310 °C (2 min), 1.5 °C/min to 315 °C, and 15 °C/min to 340 °C (2 min). The detector temperature was set to 360 °C. GC-MS analyses and identifications of individual compounds were performed on a Finnigan Trace gas chromatograph ultra coupled with a Finnigan Trace DSQ mass spectrometer (Thermo Electro Corp., Austin, TX, U.S.A.) as previously described (8–10). Solid-Phase Extraction (SPE) Lipid extracts of cereal grains were separated into fractions of free sterols/ stanols, steryl/stanyl fatty acid esters, and steryl/stanyl phenolic acid esters via SPE (Strata NH2, 55 μm, 70 Å, 1 g/6 mL, Phenomenex, Germany) as previously described (10). On-line Liquid Chromatography-Gas Chromatography (On-line LC-GC) The applied on-line LC-GC system consisted of a 1220 Infinity liquid chromatograph, which was coupled to a 7890A gas chromatograph equipped with an FID via a 1200 Infinity Series 2-position/6-port switching valve (Agilent Technologies, Waldbronn, Germany). The valve was fitted with a 200 μL sample loop. LC analyses were carried out on a 250 x 2 mm, 5 µm, Eurospher-100 Si column (Knauer, Berlin, Germany) at 27 °C using n-hexane/MTBE (96:4, v/v) as eluent for enriched dairy foods and n-hexane/MTBE/iso-propanol (96:4:0.1, v/v/v) as eluent for nuts and edible plant oils. Detection was performed with an ultraviolet (UV)-detector set to 205 nm for free sterols/stanols and steryl/stanyl esters, and set to 325 nm for trans-steryl/stanyl ferulic acid esters. LC fractions were transferred on-line by switching the valve and the evaporation of the solvent was performed via a programmable multimode inlet in the solvent vent mode. GC separations were carried out on a 30 m × 0.25 mm i.d., 0.1 µm film, 260 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.


trifluoropropylmethyl polysiloxane capillary column (Rtx-200MS, Restek GmbH, Bad Homburg, Germany). Detailed LC, GC, and interface conditions have been described elsewhere (11, 12). Identification was carried out using an on-line LC-GC-MS system. The gas chromatograph was coupled via a transfer line to an inert 5975C mass spectrometer (MS) with triple axis detector (Agilent Technologies, Waldbronn, Germany) and analyses were performed at conditions previously described (12).

Results and Discussion GC Analysis of Intact Steryl/Stanyl Fatty Acid Esters GC analysis of intact steryl/stanyl fatty acid esters is a challenge due to their structural similarities and high boiling points. Previous studies using non-polar stationary phases (e.g. DB-1 and DB-5) resulted in only insufficient separations regarding the degree of saturation of the esterified fatty acid moieties (13–15). Recently, the suitability of an intermediately polar stationary phase for the efficient separation of complex mixtures of steryl/stanyl fatty acid esters was demonstrated (8, 9, 16). The esters were separated according to the sterol/stanol moiety as well as according to the carbon number and degree of unsaturation of the esterified fatty acid moiety. The GC analysis is exemplarily shown for a commercially obtained plant stanyl fatty acid ester mixture in Figure 2. Only saturated and mono-unsaturated fatty acid esters of the same chain length eluted at the same time.

Figure 2. GC analysis of a plant stanyl fatty acid ester mixture. Peak numbering according to Table I; (IS) cholesteryl-16:0.

Taking into account the thermal instability of intact steryl/stanyl fatty acid esters during high temperature GC analysis, response factors were determined for individual esters to compensate for degradation processes (9, 16). 261 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.

The employed intermediately polar stationary phase was also shown to be effective in the detection and separation of free sterols/stanols and intact steryl/ stanyl phenolic acid esters (10). SPE-Based Approach for the Fractionation of Plant Lipids


For the separation of plant lipids into fractions containing free sterols/stanols and steryl/stanyl esters an approach based on SPE was established (10). Figure 3 illustrates the main steps of the methodology from lipid extraction to GC analysis.

Figure 3. SPE-based approach for the separation of free sterols/stanols and steryl/stanyl esters from plant lipid extracts.

The lipids were extracted from the ground plant material and further separated on an aminopropyl-modified silica gel phase. This type of phase was more efficient in the removal of triglycerides than normal silica phases. The resulting GC analyses are exemplarily shown for the fractions of free sterols/stanols, steryl/stanyl fatty acid esters, and steryl/stanyl phenolic acid esters extracted from whole corn kernels in Figure 4. On-line LC-GC-Based Approach for the Analysis of Steryl/Stanyl Fatty Acid Esters The on-line coupling of LC and GC is an efficient and elegant alternative to laborious off-line techniques such as SPE, column chromatography or thin layer chromatography. Fractionation, pre-concentration, and analysis take place in a closed and fully automated system, whereby the risks of sample loss and contamination are reduced. A schematic representation of an on-line LC-GC system with a programmable temperature vaporizer as interface for the evaporation of the solvent, which is transferred from LC to GC, is shown in Figure 5. 262 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.


Figure 4. GC analysis of (A) free sterols/stanols, (B) steryl/stanyl fatty acid esters, and (C) steryl/stanyl phenolic acid esters extracted from whole corn kernels: (1) cholesterol, (2) campesterol, (3) stigmasterol, (4) campestanol, (5) sitosterol, (6) sitostanol, (7) unknown sterol, (8) campesteryl-16:0/16:1, (9) stigmasteryl-16:0/16:1, (10) campestanyl-16:0/16:1, (11) sitosteryl-16:0/16:1, (12) sitostanyl-16:0/16:1, (13) Δ7sitosteryl-16:0/16:1, (14) campesteryl-18:0/18:1 (15) campesteryl-18:2+stigmasteryl-18:0/18:1, (16) campestanyl-18:0/18:1+stigmasteryl-18:2, (17) campestanyl-18:2, (18) Δ7campesteryl-18:2, (19) sitosteryl-18:0/18:1, (20) sitosteryl-18:2, (21) sitostanyl-18:0/18:1, (22) sitostanyl-18:2, (23) Δ7sitosteryl-18:2, (24) cis-campesteryl ferulate, (25) cis-campestanyl ferulate, (26) cis-sitosteryl ferulate, (27) cis-sitostanyl ferulate, (28) trans-campesteryl p-coumarate, (29) trans-campestanyl p-coumarate, (30) trans-sitosteryl p-coumarate, (31) trans-sitostanyl p-coumarate, (32) trans-campesteryl ferulate, (33) trans-campestanyl ferulate, (34) trans-Δ7campesteryl ferulate, (35) trans-sitosteryl ferulate, (36) trans-sitostanyl ferulate, (37) trans- Δ7sitosteryl ferulate, (38) trans-24-methylene cycloartanyl ferulate, (IS1) cholesteryl-16:0, (IS2) 5α-cholestan-3β-ol, and (IS3) trans-cholestanyl ferulate.

The lipids were fractionated on a normal silica gel phase, which has been shown to be effective for the retention of triglycerides (9, 17, 18). The fraction of steryl/stanyl fatty acid esters could then be transferred on-line to the GC, which enabled the analysis of the individual composition of the transferred LC fraction. Compared to methods commonly used for the determination of sterols/stanols and steryl/stanyl esters in enriched foods or natural foods, the advantage of the presented on-line LC-GC-based approaches is in particular the far less complex sample preparation. The work up time is strikingly decreased and compared to 263 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.


methods involving saponification and purification steps, less solvent amount is needed. Detailed information on the validation of the on-line LC-GC approach, including the limits of detection, has been provided (11, 12, 17). Figure 6 illustrates the on-line LC-GC analysis of a mixture of plant stanyl fatty acid esters.

Figure 5. Schematic presentation of an on-line LC-GC system with programmable temperature interface (transfer mode).

Figure 6. On-line LC-GC analysis of a stanyl fatty acid ester mixture extracted from an enriched cheese-based spread: (A) LC-UV chromatogram at 205 nm and (B) GC-FID chromatogram of the transferred LC fraction. Peak numbering according to Table I; (IS) cholesteryl-16:0. 264 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.

Application of the Analytical Approaches to Enriched Dairy Foods and Important Natural Sources


Investigation of Free Sterols/Stanols and Steryl/Stanyl Esters in Cereal Grains The SPE-based approach was applied to the qualitative and quantitative analysis of free sterols/stanols, steryl/stanyl fatty acid esters, and steryl/stanyl phenolic acid esters in corn, rye, wheat, and spelt (10). The methodology provided detailed data on the contents and the distributions of individual members of these compound classes. The distribution patterns of corn, particularly of steryl/stanyl fatty acid and phenolic acid esters, exhibited distinct differences compared to those of rye, wheat, and spelt. In corn, esters of sitosterol and campesterol were predominant and these sterols were mainly esterified to unsaturated fatty acids. Linoleic and oleic acid esters represented more than 90% of total steryl/stanyl fatty acid esters (Figure 7A). Sitosteryl and campesteryl esters were also predominant in rye, wheat, and spelt, but linoleic and oleic acid esters made up only approximately 50% of total esters. The other half was represented by esters of C16-fatty acids, mainly palmitic acid esters (Figure 7A).

Figure 7. Distributions of (A) steryl/stanyl fatty acid esters and (B) steryl/stanyl phenolic acid esters in cereals.

Regarding the distributions of free sterols/stanols, the observed differences were not as pronounced. Sitosterol accounted for 55-62% and was the dominating free sterol in all four investigated cereal grains, followed by either campesterol or stigmasterol. Sitostanol and campestanol made up between 10 and 16%. Within the fraction of steryl/stanyl phenolic acid esters, the trans-derivatives of sitostanyl and campestanyl ferulate were predominant. Coumaric acid esters represented less than 3.4% of total phenolic acid esters. Whereas in corn sitostanyl and sitosteryl esters were more abundant than campestanyl and campesteryl esters, the phenolic acid esters extracted from rye, wheat, and spelt showed an inverse distribution (Figure 7B). 265 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.

Table I. Analysis of Stanyl Fatty Acid Esters in Enriched Cheese-Based Spread


stanyl esters [g/100 g]a

esterified stanols [%]a,c

(1) campestanyl-16:0/16:1

0.11 ± 0.00

campestanol

29.1 ± 0.4

(2) sitostanyl-16:0/16:1

0.20 ± 0.00

sitostanol

70.5 ± 0.4

(3) campestanyl-18:0/18:1

1.00 ± 0.03

others

(4) campestanyl-18:2

0.33 ± 0.01

(5) campestanyl-18:3

0.12 ± 0.01


2.36 ± 0.05

(7) sitostanyl-18:2

0.83 ± 0.02

16:0/16:1

5.8 ± 0.1

(8) sitostanyl-18:3

0.28 ± 0.02

18:0/18:1

63.0 ± 0.4


0.08 ± 0.01

18:2

21.7 ± 0.2


0.03 ± 0.01

18:3

7.5 ± 0.4

othersb

0.03 ± 0.00

20:0/20:1

1.5 ± 0.1

total stanyl esters

5.36 ± 0.12

22:0/22:1

0.5 ± 0.1

calculated as stanolsc

3.28 ± 0.07

0.5 ± 0.0

esterified fatty acids [%]a,c

a Values represent the mean × standard deviation of a triplicate analysis. b Calculated with a response factor of 1, relative to cholesteryl-16:0. c Calculated on the basis of intact stanyl esters.

Investigation of Stanyl Fatty Acid Esters in Fat-Based Enriched Dairy Foods On-line LC-GC was successfully applied to the analysis of intact steryl/stanyl esters in enriched dairy products with substantial amounts of protein and fat (11). The samples were subjected to an acid digestion step with hydrochloric acid to release the lipids from the protein matrix, followed by the extraction of the lipids using a mixture of n-hexane and MTBE. The lipid extracts were further subjected to on-line LC-GC analysis for the determination of the contents and compositions of the added stanyl fatty acid esters. Possibly interfering neutral lipids such as triglycerides could be effectively removed using a normal silica gel phase as stationary LC-phase and n-hexane/MTBE (96:4, v/v) as eluent. Hence, no further purification step was needed and the extracts could directly be analyzed with the on-line LC-GC system. Table I shows qualitative and quantitative data on total and individual stanyl fatty acid esters obtained by the investigation of an enriched cheese-based spread. Oleic acid esters were predominant, followed by linoleic, linolenic, and palmitic acid esters. The determined amounts were in agreement with the declaration on the package (3.3 g stanols/100 g), and the calculated profile of the esterified fatty acids indicates rapeseed oil being the source of the fatty acid mixture used for the esterification of the stanols. 266 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.


Investigation of Free Sterols/Stanols, Steryl/Stanyl Esters, and Other Minor Lipids in Edible Plant Oils and Nuts The sample preparation for the analysis of free sterols/stanols, steryl/stanyl esters, and other minor lipids in edible plant oils and nut lipids only required a silylation of the oils. Owing to the silylation step, the trimethyl silyl (TMS)derivatives of free sterols/stanols and the steryl/stanyl fatty acid esters as well as the TMS-derivatives of other minor lipids such as tocopherols and free fatty acids exhibited similar polarities and thus eluted at the same time in a merged peak under the employed LC conditions; this enabled the transfer in a single fraction to the GC (12). Trans-derivatives of steryl/stanyl ferulic acid esters could be analyzed via a second transfer. The simultaneous analysis of free fatty acids, tocopherols, free sterols, and steryl fatty acid esters by on-line LC-GC is exemplarily shown for rapeseed oil in Figure 8.

Figure 8. On-line LC-GC analysis of free fatty acids, tocopherols, free sterols, and steryl fatty acid esters in rapeseed oil: (A) LC-UV chromatogram at 205 nm and (B) GC-FID chromatogram of transferred LC fraction: (1) δ-tocopherol, (2) γ-tocopherol, (3) cholesterol, (4) α-tocopherol, (5) brassicasterol, (6) campesterol, (7) stigmasterol, (8) sitosterol, (9) brassicasteryl-16:0/16:1, (10) campesteryl-16:0/16:1, (11) sitosteryl-16:0/16:1, (12) brassicasteryl-18:0/18:1, (13) brassicasteryl-18:2, (14) brassicasteryl-18:3, (15) campesteryl-18:0/18:1, (16) campesteryl-18:2, (17) campesteryl-18:3, (18) sitosteryl-18:0/18:1, (19) sitosteryl-18:2, (20) sitosteryl-18:3, (IS1) 5α-cholestan-3β-ol, and (IS2) cholesteryl-16:0. The analysis of several commercially available oils revealed corn germ oil and rapeseed oil as the richest sources of, in particular, steryl/stanyl fatty acid esters. The average total amounts of steryl/stanyl fatty acid esters ranged from 0.07 to 0.96 mg/100 mg oil; those of free sterols/stanols from 0.14 to 2.34 mg/100 mg oil (Figure 9). Ferulic acid esters could only be detected in corn germ oil, 267 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.


accounting for 0.1 mg/100 mg oil. The majority of the sterols/stanols in most of the investigated plant oils occurred in form of their fatty acid esters, except for the native sunflower oils, safflower oil, soybean oil and olive oils, where the amounts of free sterols/stanols and steryl/stanyl fatty acid esters were either equal or free sterols/stanols were predominant. Regarding the distribution patterns of free sterols/stanols and steryl/stanyl esters, considerable differences were observed between the various oils. The applied on-line LC-GC-based approach enabled the fast and robust analysis of these lipid compounds, which could thus be a useful tool for authenticity assessments of plant oils.

Figure 9. Mean total contents of free sterols/stanols and steryl esters in edible plant oils. Additionally, three batches of ten different commercially important nut types were studied regarding their contents and compositions of free sterols/stanols and steryl/stanyl fatty acid esters (Figure 10).

Figure 10. Mean total contents of free sterols/stanols and steryl/stanyl esters in oils extracted from nuts. 268 In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.

The main part of the sterols/stanols in the investigated nuts occurred in free form, followed by fatty acid esters; trans-steryl/stanyl ferulic acid esters could not be detected. The total amounts of steryl/stanyl fatty acid esters in the extracted nut oils ranged from 0.02 to 0.28 mg/100 mg, those of free sterols/stanols from 0.12 to 0.37 mg/100 mg; they were both by far the highest in pistachios and pine nuts (12).


Conclusion Capillary gas chromatography on an intermediately polar stationary phase was shown to be suitable for the detection and separation of individual free sterols/stanols and intact steryl/stanyl esters. On this basis, analytical approaches for the qualitative and quantitative analysis of free sterols/stanols and steryl/stanyl esters were established. The authentication of steryl/stanyl fatty acid esters in enriched fat-based dairy foods can be performed via a combination of acid digestion, lipid extraction, and on-line LC-GC analysis. For the determination of naturally occurring free sterols/stanols and steryl/stanyl esters, an approach based on solid-phase extraction and GC analysis was developed; the applicability was shown for the investigation of cereal grains. The suitability of on-line LC-GC for the analysis of free sterols/stanols and steryl/stanyl esters in plant lipids was demonstrated using edible plant oils and nuts as examples. The established methodologies show distinct advantages compared to methods commonly used for the determination of sterols/stanols and steryl/stanyl esters in enriched as well as natural foods. They allow the simultaneous analysis of various sterol/stanol substance classes, and especially the achieved GC separation enables a detailed qualitative and quantitative analysis of individual intact steryl/stanyl esters. Thus, the presented methodologies can be applied as useful tools for the analytical characterization of functional foods and of naturally occurring bioactive food constituents.

References 1. 2.

3. 4. 5. 6. 7.

Piironen, V.; Lindsay, D. G.; Miettinen, T. A.; Toivo, J.; Lampi, A.-M. J. Sci. Food Agric. 2000, 80, 939–966. Demonty, I.; Ras, R. T.; van der Knaap, H. C. M.; Duchateau, G. S. M. J. E.; Meijer, L.; Zock, P. L.; Geleijnse, J. M.; Trautwein, E. A. J. Nutr. 2009, 139, 271–284. Plat, J.; Mensink, R. P. Am. J. Cardiol. 2005, 96, 15D–22D. Katan Martijn, B.; Grundy Scott, M.; Jones, P.; Law, M.; Miettinen, T.; Paoletti, R. Mayo Clin. Proc. 2003, 78, 965–78. SCF (Scientific Committee on Food) SCF/CS/NF/DOS/1 FINAL. 2000. SCF (Scientific Committee on Food) SCF/CS/NF/DOS/15 ADD 2 FINAL. 2003. SCF (Scientific Committee on Food) SCF/CS/NF/DOS/23 ADD 2 FINAL. 2003. 269

In Instrumental Methods for the Analysis and Identification of Bioactive Molecules; Jayprakasha, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.

8. 9. 10. 11.


12. 13. 14. 15. 16.

17. 18.

Barnsteiner, A.; Esche, R.; di Gianvito, A.; Chiavaro, E.; Schmid, W.; Engel, K.-H. Food Control 2012, 27, 275–283. Barnsteiner, A.; Lubinus, T.; di Gianvito, A.; Schmid, W.; Engel, K.-H. J. Agric. Food Chem. 2011, 59, 5204–5214. Esche, R.; Barnsteiner, A.; Scholz, B.; Engel, K.-H. J. Agric. Food Chem. 2012, 60, 5330–5339. Esche, R.; Carcelli, A.; Barnsteiner, A.; Sforza, S.; Engel, K.-H. Eur. Food Res. Technol. 2013, 236, 999–1007. Esche, R.; Müller, L.; Engel, K.-H. J. Agric. Food Chem. 2013, 61, 11636–11644. Evershed, R. P.; Male, V. L.; Goad, L. J. J. Chromatogr., A 1987, 400, 187–205. Gunawan, S.; Melwita, E.; Ju, Y.-H. Food Chem. 2010, 121, 752–757. Kamm, W.; Dionisi, F.; Fay, L. B.; Hischenhuber, C.; Schmarr, H. G.; Engel, K. H. J. Chromatogr., A 2001, 918, 341–349. Engel, K.-H.; Barnsteiner, A. In Progress in Authentication of Food and Wine; Ebeler, S. E., Takeoka, G. R., Winterhalter, P., Eds.; ACS Symposium Series 1081; American Chemical Society: Washington, DC, 2011; pp 177−187. Esche, R.; Scholz, B.; Engel, K.-H. J. Agric. Food Chem. 2013, 61, 10932–10939. Grob, K.; Kaelin, I.; Artho, A. J. High Resolut. Chromatogr. 1991, 14, 373–376.

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GC and On-line LC-GC - American Chemical Society

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