C0532011019

IOSR Journal Of Pharmacy
(e)-ISSN: 2250-3013, (p)-ISSN: 2319-4219
www.iosrphr.org Volume 5, Issue 3 (March 2015), PP. 11-19
11
Validation and Application of a Gas Chromatographic Method
for the Determination of Fatty Acids and Sterols in the Total Diet
Vesna Kostik
Institute of Public Health of Republic of Macedonia, 50 Divizija No. 6, 1000 Skopje, Republic of Macedonia
ABSTRACT : Fatty acids (FAs) and sterols represent the most important fraction of edible oils and fats,
which play a significant role in human nutrition and health. A gas chromatographic method for the
identification and quantification of FAs and sterols based on the extraction of lipids by accelerated solvent
extraction (ASE), saponification of the obtained extract, and direct determination of the isolated FAs and sterols
without derivatization, was developed and validated. The proposed method was accurate and precise (mean
recovery in the range 80.8% to 98.2%, precision with RSD < 8.6%; LOQ < 0.86 mg/kg; measurement
uncertainty < 14.8%). This method was applied for the determination of the most abundant FAs and sterols in
the total diet. The obtained results showed that the most abundant FA in the total diet is monounsaturated C18:1
with 34.5% of the total FAs. The lowest presence was observed for the short chain FAs: C4:0 – C12:0 (6.4%).
Cholesterol was found to be the most abundant sterol in the diet (57.5%). The total amount of the plant sterols
in the diet was found to be 42.5%.
KEYWORDS: gas chromatography, fatty acid, sterol.
I. INTRODUCTION
Edible oils and fats are one of the major components of human diets, comprising as much as 25% of
average caloric intake. They play important functional and sensory roles in food products and they act as
carriers of fat-soluble vitamins (A, D, E, and K). They also provide an essential linoleic and linolenic acid,
responsible for growth [1]. Edible oils and fats are biological mixtures of plant origin consisting of ester
mixtures derived from glycerol with chain of fatty acids [2]. Both the physical and the chemical characteristics
of oils and fats are greatly influenced by the kind and proportion of the fatty acids (FAs) on the triacylglycerol
[3, 4]. FAs can be classified in classes as saturated, mono-unsaturated (MUFA) and polyunsaturated (PUFA)
fatty acids. The predominant fatty acids present in the diet are saturated and unsaturated compounds with
straight aliphatic chains. An even number of carbon atoms, from 16 to 18, with a single carboxyl group, is the
most common [5, 6]. High levels of saturated FAs in the diet are frequently considered to have influence in
increasing the concentration of low density lipoproteins (LDL), affecting the ratio of LDL to HDL (high density
lipoproteins), promoting clothing and vascular smooth muscle proliferation [7, 8]. Diet with increasing intake of
linoleic and linolenic acids increase HDL-cholesterol and decreases LDL-cholesterol, while higher intake of
oleic acid decreases LDL-cholesterol, but does not affect HDL cholesterol levels [9].
Therefore, the determination of FAs is an essential requirement in testing food material [10].Gas
chromatography (GC) coupled with mass spectrometer (MS) or flame ionization detector (FID) is the most
widely used technique for the determination of FAs [11-13]. Commonly, the most methodologies used for
determining FAs are lipid extraction followed by conversion of the FAs into corresponding methyl esters
(FAMEs) [14, 15]. Such methodologies are usually used for preparing FAMEs from lipids either by basic
hydrolysis followed by methylation of the free FAs (FFAs) or by trans-esterification of lipids using acid or base
catalyzed as rapid and simple methods [14,16].
The presence of non-lipids at the moment of the derivatization process may interfere with lipids
resulting to potential errors and high variable profiles [17].
Beside fatty acids, the most abundant components present in the edible oils and fats are sterols.
Cholesterol, sitosterol and stigmasterol are polycyclic steroid compounds with similar chemical structure.
Cholesterol is a typical animal sterol, e.g. its content in milk fat is 95–98%, stigmasterol and sitosterol are
referred to as phytosterols [18].
The determination of sterols in the total diet is carried out for the following objectives: to measure the
total cholesterol content to obtain nutritional information and to detect the presence of vegetable fats [18,19].
The most appropriate and frequently used method for the determination of total sterols content in foods
is the direct saponification, followed by the extraction of the unsaponificable residue into the nonpolar solvent,
derivatization of the isolated sterols and final gas chromatographic detection [20]. The aim of our study was to
develop and validate simple, reliable and accurate GC method for determination of the most abundant FAs and

Valdiation and application of a gas chromatographic method for the determination of fatty…
12
sterols in the total diet without derivatization of the isolated analytes. The method was applied for determination
of FAs and sterols in the total diet (28 samples).
.
II. MATERIALS AND METHODS
2.1. Instrumentation
The chromatographic analysis (GC) were performed on a Shimadzu 2010 GC system equipped with
flame-ionization detector (FID), and auto injector (AOC- 20i), and the Chrom- Solution software. The sample
extraction was performed in a DIONEX Accelerated Solvent extractor, ASE-100 (USA). Stainless steel
extraction cells (34 mL) were used for the extraction. Helium (purity 99.999 %) was used as a purge gas. The
sample saponification was performed with reflux condenser.
2.2. Reagents and standards
All chemicals and solvent were a special grade for gas chromatography. N-hexane, 96% (V/V) ethanol,
absolute ethanol and chloroform were obtained from Merck (Darmstadt, Germany). Anhydrous sodium
sulphate, (prepared 3 hours at 650 0
C), was purchased from Sigma-Aldrich/Fluka/Riedel-de-Haen (Zwijndrecht,
The Netherlands). The moisture absorbing polymer (MAP) and diatomaceous earth (DE) were purchased from
Thermo Fisher Scientific, Waltham, MA (USA). Water was deionized then distilled from glass apparatus.
Potassium hydroxide analytical grade was obtained from Merck (Darmstadt, Germany)
The analytical standards: butyric acid (C4:0) purity ≥98.5% GC, caproic acid (C6:0) purity ≥99.0% GC,
caprylic acid (C8:0) purity ≥98.5% GC, capric acid (C10:0) purity ≥99.5% GC, lauric acid (C12:0) purity ≥99.0%
GC, myristic acid (C14:0) purity ≥98.5% GC, palmitic acid (C16:0) purity ≥99.0% GC, palmitoleic acid (C16:1)
purity ≥98.5% GC, stearic acid (C18:0) purity ≥99.0% GC, oleic acid (C18:1) purity ≥98.5% GC, linoleic acid
(C18:2) purity ≥98.5%, linolenic acid (C18:3) purity ≥98.5%, cholesterol purity ≥98.5% GC, stigmasterol purity
≥98.5% GC, β- sitosterol purity ≥98.0% GC, campesterol purity ≥98.5% GC and 5-α-cholestane purity ≥98.0%
GC were obtained from Sigma-Aldrich/Fluka/Riedel-de-Haen (Zwijndrecht, The Netherlands). Blank samples
of dietary non fatty supplements used for fortification were purchased from the local market.
2.3. Preparation of the standard solutions
2.3.1. Preparation of FAs standard solutions
Stock solutions of individual fatty acid (FA) standards were prepared in n – hexane at 1000 mg/L and
were stored in a refrigerator at -20 0
C. Standard working solution mixture (i) was prepared by transferring 10
mL of each individual stock standard solution in a 50 mL volumetric flask and diluting with n – hexane to a
concentration of 200 mg/L. Stock standard solutions of individual FA and standard working solution mixture (i)
was used for the preparation of chromatographic standard solutions (ii) with different FA concentrations: 10
mg/L – 100 mg/L, i.e. 10 mg/L; 25 mg/L; 50 mg/L; 75 mg/L and 100 mg/L. Chromatographic standard
solutions were prepared in n-hexane. Stock solutions and standard working solution mixture (i) were used for
the fortification of the blank samples.
2.3.2. Preparation of sterols standard solutions
Stock solutions of individual sterols were prepared in chloroform at 1000 mg/L and stored in a
refrigerator at -200
C. Standard working solution mixture (i) was prepared by transferring 5 mL of each
individual stock standard solution in a 50 mL volumetric flask and diluting with chloroform to a concentration
of 100 mg/L. Standard working solution mixture (ii) was prepared by transferring 5 mL of standard working
solution mixture (i) to a 50 mL volumetric flask and diluting with chloroform to a concentration of 10 mg/L.
Standard working solution mixture (i) was used for the preparation of the chromatographic standard solutions
with different stigmasterol and campesterol concentrations: 1 mg/L – 10 mg/L, i.e. 1.0 mg/L; 2.5 mg/L; 5 mg/L;
7.5 mg/L and 10 mg/L. Stock solutions of cholesterol and β sitosterol were used for the preparation of the
chromatographic standard solutions with cholesterol and β sitosterol concentrations ranging from 10 mg/L to
100 mg/L, i.e. 10 mg/L; 25 mg/L; 50 mg/L; 75 mg/L and 100 mg/L. Chromatographic standard solutions were
prepared in chloroform. Stock solutions and standard working solution mixture (i) and mixture (ii) were used for
the fortification of blank samples. Internal standard stock solution of 5-α-cholestane was prepared in chloroform
at a 1000 mg/L. Working internal standard solution was prepared by transferring 100 μL of internal standard
stock solution (IS) in a 10 mL volumetric flask and diluting with chloroform to a concentration of 10 mg/L. 1
mL of internal standard solution (10 mg/L) was added into the sample prior the extraction.

13
2.4. Sample preparation
An aliquot of 10 g of homogenized sample was mixed with 2 g of MAP and 2 g of DE. The extraction
cell was filled with the homogenized mixture, and placed in the ASE, which worked out under the conditions
shown in Table 1.
Table 1. ASE operating conditions
Solvent n-hexane
Temperature 100 0
C
Pressure 1500 psi
Static time 5 min
Flush volume 60 %
Purge time 140 sec
Static Cycle 1
The obtained extract (15 mL) was evaporated under the stream of nitrogen to dryness. The residue was
dissolved in 30 mL solution of KOH in absolute ethanol (0.5 mol/L). The saponification was performed using
reflux condenser within 30 min. The solution obtained after the saponification was quantitatively transferred
into a 50 mL volumetric flask and filled with absolute ethanol up to the mark. The solution was used for the
further analysis.
2.4.1. Extraction of FAs
5 mL of the solution was transferred to a 100 mL separatory funnel. 0.1 mL 25% (V/V) HCl was added
together with 25 mL of n-hexane and 25 mL of distilled water. The mixture was vigorously shaken for 5 min and
then allowed to stand for 15 min. The water portion was discarded. The hexane layer was collected through the
anhydrous sodium sulphate and removed by a rotary evaporator at 40 0
C. The residue was dissolved in n –
hexane and adjusted to the volume of 10 mL with the same solvent. The sample solution was used for GC-FID
determination.
2.4.2. Extraction of sterols
45 mL of the solution was transferred to a 250 mL separatory funnel together with 100 mL of n-hexane
and 50 mL of mixture (96%, V/V) ethanol: distilled water (1:1). The mixture was vigorously shaken for 5 min
and then allowed to stand for 15 min. The water portion was discarded. The hexane layer was collected through
the anhydrous sodium sulphate and removed by a rotary evaporator at 40 0
C. The residue was dissolved in
chloroform and adjusted to the volume of 1 mL with the same solvent. The sample solution was used for GC-
FID determination.
2.5. GC determination
Separation of FAs was achieved on a fused silica HP-FFAF capillary column (25 m x 0.32 mm i.d. x
0.52 μm film thickness), supplied by Agilent (USA). Operating conditions were as follows: injector port
temperature, 230 0
C; injection volume, 2 μL in split modе (1:10); detector temperature, 260 0
C (make up gas -
nitrogen flow 27.5 mL/min); nitrogen as carrier gas at flow rate at 1.5 mL0
C /min; oven temperature
programme, 180 0
C (1 min) increased with the rate of 2 0
C/min to 200 0
C and held for 19 min. The total run time
of the chromatographic analysis was 30 min. The column equilibration time was 3 min.
Sterols separation was achieved on a fused silica HP-FFAF capillary column (25 m x 0.32 mm i.d. x
0.52 μm film thickness), supplied by Agilent (USA). Operating conditions were as follows: injector port
temperature, 230 0
C; injection volume, 2 μL in split mode (1:3); detector temperature, 300 0
C (make up gas -
nitrogen flow 27.5 mL/min); nitrogen as carrier gas at flow rate at 1.5 mL0
C /min; oven temperature was set at
260 0
C and held for 40 min. The column equilibration time was 3 min.
The proposed method was validated in respect to the recovery, linearity, precision expressed as within
day repeatability and between day reproducibility of retention time and peak area, stability, limit of detection
(LOQ), limit of quantification (LOQ) and measurement uncertainty (Ux).
III. RESULTS AND DISCUSSION
3.1. Method optimization
In order to obtain the best separation for all tested FAs and sterols, a series of preliminary
investigations with capillary GC columns with a different polarity of the stationary phase were tested. The
optimum separation of components of interest was achieved when capillary column with a strong polarity
(Polyethylene glycol-TPA modified) and optimized temperature programmes are used (Fig. 1).

14
Representative chromatograms for FAs and sterols separation with GC – FID method are shown in Fig.
1 and Fig. 2, respectively.
Figure 1. Chromatogram of standard solution of C16:0 (1), C18:0 (2), C18:1 (3), C18:2 (4) and C18:3 (5), at 10 mg/L
Figure 2. Chromatogram of the sterols extract from total diet sample [1. 5 α-Cholestan (I.S.); 2. Cholesterol; 3.
Stigmasterol; 4. Campesterol; 5. β - Sitosterol]
In our research accelerated solvent extraction (ASE) technique was used for samples extraction. The
elevated pressure increases the boiling temperature of the solvent and extraction can occur at relatively high
temperatures, which leads to faster extractions [21, 22]. The extraction process is therefore significantly faster
than traditional methods such as Soxhlet extraction or supercritical fluid extraction [23]. In a study conducted by
Omar and Salimon [20] the total time required for extraction of the lipids from the sample with Soxhlet
extraction was approximately 3 hours. Soxhlet extraction and SFE are also used as techniques for the isolation
of lipids prior to the determination of sterols [24]. In our investigations due to the use of elevated temperature
and pressure in the extraction cell the extraction time was approximately 15 min.
Up to our knowledge, no previous data was reported in the literature for direct determination of FAs
and sterols by GC. In our study, for the first time we introduce an analytical method for direct determination of
free FAs and sterols, avoiding the derivatization step and therefore the possible errors which could be due to this
demanding procedure.
3.2. Method Validation
3.2.1. Recovery, Precision and Linearity
The recovery, precision and linearity of all the FAs and sterols was determined using fortified samples
of dietary non fatty supplements which were previously tested on the presence of FAs and sterols. In each case,
5 replicates each at 5 levels were fortified into the samples. The samples (10 g) were fortified before the
extraction to yield 10 mg/kg; 25 mg/kg; 50 mg/kg; 75 mg/kg and 100 mg/kg, respectively (C4:0, C6:0, C8:0, C10:0,
C16:0, C16:1, C18:0, C18:1, C18:2, C18:3, cholesterol and β - Sitosterol); and to yield 1.0 mg/kg; 2.5 mg/kg; 5 mg/kg; 7.5

15
mg/kg and 10 mg/kg, respectively (stigmasterol and campesterol). Gas tight glass syringes (10 µL, 50 µL and
100 µL) were use for the addition of spiking mixtures.
The recovery, precision and linearity of the tested method for the determination of FAs and sterols are
presented in Table 2 and Table 3, respectively.
From the obtained results, it can be noticed that the proposed method is accurate and precise enough for
the determination of all the tested FAs and sterols. The obtained values for multiple correlation coefficients,
ranged from 0.9904 to 0.9982, indicated that the method has a good linearity for all the tested compounds. High
analytical recoveries ranging from 80.8% to 98.2% were obtained for FAs determination, as well as, for sterols
determination (82.7% - 91.0%).
Table 2. Statistical data for mean recovery, precision data and linearity of the method for the
determination of FAs
FA
Fortification level
(mg/kg)
FA found - mean value
(mg/kg ± SD)
Recovery (%), n=5 RSD (%) Regression equation
C4:0
10 8.98 ± 0.55 89.8 6.2
y = 0.9912x – 2.2561
R2
= 0.9987
25 22.4 ± 1.2 86.2 5.3
50 45.6 ± 2.4 91.1 5.2
75 71.3 ± 4.0 95.1 5.6
100 98.2 ± 4.7 98.2 4.8
C6:0
10 9.12 0.23 91.2 2.5
y = 0.9832x – 1.6415
R2
= 0.9982
25 23.5 ± 1.05 94.0 4.5
50 44.8 ± 2.5 89.6 5.6
75 72.5 ± 3.4 96.7 4.7
100 97.5 ± 4.2 97.5 4.3
C8:0
10 8.79 ± 0.61 87.9 6.9
y = 0.969x – 2.7665
R2
= 0.9954
25 21.8 ± 1.2 87.2 5.5
50 41.5 ± 2.1 83.0 5.1
75 70.5 ± 2.3 94.0 3.3
100 95.5 ± 4.4 95.5 4.6
C10:0
10 8.65 ± 0.54 86.5 6.2
y = 0.9525x – 3.0378
R2
= 0.9951
25 20.5 ± 1.1 82.0 5.4
50 40.5 ± 1.9 81.0 4.7
75 69.6 ± 2.9 92.8 4.2
100 93.2 ± 5.2 93.2 5.6
C12:0
10 8.3 ± 0.40 83.0 4.8
y = 0.9136x – 1.4293
R2
= 0.9983
25 21.4 ± 1.4 85.6 6.8
50 42.3 ± 2.5 84.6 5.9
75 68.9 ± 3.1 91.9 4.5
100 89.5 ± 5.9 89.5 6.6
C14:0
10 8.95 ± 0.54 89.5 6.8
y = 0.9089x –1.9915
R2
= 0.9982
25 20.5 ± 1.6 82.3 7.8
50 41.4 ± 2.8 82.8 6.7
75 67.9 ± 3.6 90.5 5.3
100 88.6 ± 5.5 88.6 6.2
C16:0
10 8.15 ± 0.6 81.5 7.4
y = 0.9273x –1.8305
R2
= 0.9987
25 20.8 ± 1.8 83.2 8.6
50 43.2 ± 2.5 86.4 5.8
75 69.5 ± 3.6 92.7 5.2
100 90.3 ± 5.2 90.3 5.7
C16:1
10 8.8 ± 0.43 88.0 5.5
y = 0.9411x –2.4756
R2
= 0.9956
25 22.3 ± 1.3 89.2 5.8
50 40.5 ± 2.1 81.6 5.2
75 69.4 ± 2.9 92.5 4.2
100 92.3 ± 4.8 92.3 5.2
C18:0
10 8.6 ± 0.45 86.0 5.9
y = 0.952x –2.5634
R2
= 0.9926
25 23.4 ± 1.6 93.6 6.8
50 39.8 ± 2.5 89.6 6.3
75 70.5 ± 3.2 94.0 4.5
100 93.4 ± 5.1 93.4 5.5
C18:1
10 8.3 ± 0.40 83.0 5.4
y = 0.9464x – 3.3537
R2
= 0.9906
25 20.2 ± 1.4 80.8 6.9
50 39.6 ± 2.8 89.2 7.1
75 72.4 ± 3.0 96.5 4.1
100 89.8 ± 4.9 89.8 5.5
C18:2
10 8.1 ± 0.50 81.0 7.0
y = 0.9297x – 3.2634
R2
= 0.9912
25 20.5 ± 1.3 82.0 6.3
50 38.5 ± 2.1 82.5 5.4

16
FA
Fortification level
(mg/kg)
(mg/kg ± SD)
Recovery (%), n=5 RSD (%) Regression equation
75 70.5 ± 3.2 94.0 4.5
100 88.8 ± 4.7 88.8 5.3
C18:3
10 8.34 ± 0.30 83.4 4.3
y = 0.9192x – 3.5561
R2
= 0.9904
25 19.8 ± 1.2 84.3 6.1
50 37.5 ± 2.4 85.0 6.4
75 69.5 ± 2.9 92.7 4.2
100 87.5 ± 3.6 87.5 4.1
Table 3. Statistical data for mean recovery, precision data and linearity of the method for the determination of
sterols
FA
Fortification level
(mg/kg)
(mg/kg ± SD)
Recovery (%),
n=5
RSD (%) Regression equation
Cholesterol
10 8.3 ± 0.42 83.0 6.3
y = 0.8744x – 3.9683
R2
= 0.9953
25 18.6 ± 1.6 84.4 8.6
50 36.5 ± 2.7 83.0 7.4
75 60.5 ± 3.6 82.9 5.9
100 85.6 ± 4.5 85.6 5.2
β - Sitosterol
10 8.93 ± 0.30 89.3 4.3
y = 0.8421x – 2.4683
R2
= 0.9975
25 19.2 ± 1.4 86.8 7.3
50 37.6 ± 2.4 87.5 6.4
75 59.5 ± 3.4 86.3 5.7
100 83.4 ± 4.7 83.4 5.6
Campesterol
1.0 0.863 ± 0.03 86.3 4.3
y = 0.9274x – 0.3024
R2
= 0.9929
2.5 2.1 ± 0.15 84.5 7.1
5.0 3.9 ± 0.20 85.0 5.1
7.5 7.0 ± 0.30 83.7 4.3
10 8.9 ± 0.40 89.0 4.5
Stigmasterol
1.0 0.837± 0.04 83.7 5.5
y = 0.943x – 0.3378
R2
= 0.9975
2.5 2.0 ± 0.15 85.4 7.5
5.0 4.1 ± 0.22 83.6 5.4
7.5 6.9 ± 0.35 92.0 5.1
10 9.1 ± 0.30 91.0 3.3
3.2.2. Repeatability and Reproducibility
The within day repeatability of our method was determined by performing the analysis of 10 blank
samples fortified with FAs at 50 mg/g and sterols at 10 mg/kg, respectively. After the isolation of the lipid
fraction, saponification of the extracts and extraction of FAs and sterols, obtained extracts were analyzed by GC-
FID within the same day under the chromatographic conditions described in the section 2.5. The between day
reproducibility of the method was determined by performing the analysis of the extracts obtained from 5
fortified blank sample (50 mg/g of FAs, 10 mg/L of sterols). After the isolation of the lipid fraction,
saponification of the extracts and extraction of FAs and sterols, obtained extracts were analyzed by GC-FID
within the same day under the chromatographic conditions described in the section 2.5. The calculated RSD
values for the retention time (tR), the peak areas, within day repeatability and between day reproducibility are
shown in Table 4 and Table 5, respectively.
The calculated RSD values for within day repeatability in the case of FAs determination for tR values
ranged from 0.112% to 0.190%, whereas for peak areas RSD values ranged from 3.03% to 4.61%, indicating
good precision of the tR, as well as of the peak area, within the same day. Good precision of the tR, (RSD from
0.165% to 0.196%) and the peak area (RSD from 3.23% to 4.95%) were also observed within different days
(between day reproducibility).
The calculated RSD values for within day repeatability in the case of sterols determination for tR values
ranged from 0.182% to 0.192%, whereas for peak areas RSD values ranged from 3.82% to 4.27%, indicating
good precision of the tR, as well as of the peak area, within the same day. Good precision of the tR, (RSD from
0.196% to 0.205%) and the peak area (RSD from 4.11% to 4.94%) were also observed within different days
(between day reproducibility).

17
Table 4. Statistical data for within day repeatability and between day reproducibility for the determination of
FAs
FA tR /min
Within day Repeatability (RSD,
%); n=10
Between day Reproducibility (RSD,
%); n=25
C4:0 4.872 0.112 4.17 0.165 4.11
C6:0 6.782 0.162 3.91 0.179 3.23
C8:0 8.450 0.190 3.40 0.198 3.78
C10:0 9.454 0.110 3.73 0.184 3.94
C12:0 12.345 0.121 4.61 0.194 4.97
C14:0 14.190 0.103 3.23 0.172 4.56
C16:0 15.104 0.105 3.52 0.198 4.46
C16:1 21.146 0.163 3.29 0.189 4.95
C18:0 21.890 0.142 3.79 0.177 4.65
C18:1 22.267 0.164 3.85 0.180 4.90
C18:2 23.467 0.161 3.21 0.192 4.34
C18:3 25.796 0.176 3.03 0.196 4.95
Table 5. Statistical data for within day repeatability and between day reproducibility for the determination of
sterols
FA tR /min
Within day Repeatability (RSD,
%); n=10
Between day Reproducibility (RSD,
%); n=25
Cholesterol 15.163 0.192 4.27 0.205 4.11
Stigmasterol 18.893 0.182 3.97 0.219 4.23
Campesterol 20.983 0.195 3.82 0.238 4.78
β -Sitosterol 29.124 0.185 3.93 0.196 4.94
3.2.3. Stability
Stock standard solutions and working standard solutions were found to be stable for at least 3 months,
respectively, when stored at -20 0
C. Moreover, the stability of a fortified blank sample at a concentration of 50
mg/g kept in the auto injector for 24 hours was assayed, and differences of < 5.0 % were obtained.
3.2.4. Limit of Detection, Limit of Quantification, Measurement Uncertainty
The limit of detection (LOD) and the limit of quantification (LOQ) were calculated according to the
formulas LOD = 3.3 · SD/slope and LOQ = 10 · SD/slope [25].
Table 6. Statistical data for LOD, LOQ, Ux for FAs and sterols
Compound LOD (mg/kg) LOQ (mg/kg) Ux (%)
C4:0 0.22 0.73 12.5
C6:0 0.18 0.59 14.8
C8:0 0.16 0.53 11.3
C10:0 0.19 0.63 11.9
C12:0 0.21 0.69 10.8
C14:0 0.15 0.49 9.6
C16:0 0.12 0.40 11.6
C16:1 0.15 0.49 10.5
C18:0 0.16 0.53 8.5
C18:1 0.10 0.33 9.5
C18:2 0.11 0.36 11.2
C18:3 0.12 0.40 9.8
Cholesterol 0.18 0.59 13.7
Stigmasterol 0.25 0.82 13.4
Campesterol 0.22 0.73 11.2
β -Sitosterol 0.26 0.86 12.4
The values for measurement uncertainty (Ux) were calculated according to a Eurachem/CITAC Guide
[26]. The obtained values showed in Table 5, ranged from 8.5% to 14.8%.
3.3. Application of the method
In order to demonstrate the applicability of the proposed method for the determination of FAs and
sterols in the total diet, the 2 week experiment was conducted. Sampling was done according to the
methodology of Thomas et al. [27]. The results of the investigation are presented in Fig. 3 and Fig. 4,
respectively.

18
Figure 3. Distribution of the most abundant FAs in the total diet
Figure 4. Distribution of the most abundant sterols in the total diet
As can be seen from the obtained results, the most abundant FA in the total diet is monounsaturated
C18:1 with 34.5% of the total FAs. The lowest presence was observed for the short chain FAs: C4:0 – C12:0 (6.4%).
Cholesterol was found to be the most abundant sterol in the diet (57.5%). The total amount of the plant sterols in
the diet was found to be 42.5%.
IV. CONCLUSION
A reliable, accurate and precise GC-FID method, after extraction of the samples with ASE,
saponification of the lipid fraction and extraction of the FAs and sterols with n-hexane from the same solution
was developed and validated. With the usage of ASE technique, extraction of lipids from the samples was
performed within 15 minutes. The proposed extraction procedure of the FAs and sterols enables the single step
requiring the same organic solvent. The proposed method, reduce the amount of co- extractible lipids and
disables the appearance of the background peaks in the chromatogram, above signal to noise ratio of 3, at the
retention times of targeted compounds. FID was chosen for this analysis due to its selectivity for the compounds
of interest. Validation parameters obtained for determination of the FAs and sterols in the total diet demonstrate
that the developed analytical method meets the method performance acceptability criteria (mean recovery in the
range 80.8% to 98.2%, precision with RSD < 8.6%; LOQ < 0.86 mg/kg; measurement uncertainty < 14.8%).
REFERENCES
[1]. O.O.Fasina , C.H.M.Hallman, and C. Clementsa, Predicting Temperature-Dependence Viscosity of Vegetable Oils from Fatty Acid
Composition, JAOCS, 83(10), 2006, 899-903.
[2]. M.A.D Eqbql, A.S Halimah, M.K. Abdulah, and M.K Zalifah, Fatty acid composition of four different vegetable oils (red palm
olein,corn oil and coconut oil) by gas chromatography, IPCBE, 14, 2011, 31-34.
[3]. S. P J. N. Senanayake, and F.Shahidi, Structured lipids: acydolysis of gamma-linolenic acid rich-oils with n-3 polyunsaturated fatty
acids, Journal of food lipids, 4, 2002, 309-323.
[4]. A. J. St. Angelo, Lipid oxidation in foods,Critical review in Food Science and Nutrition 36 (3), 1996, 175-224.

19
[5]. M. Daniewski , B. Jacorzynski , A. Filipek, J. Balas , M. Pawlizka, and E. Mielniczuk (2003), Fatty acid content in selected edible
oils, Roczniki-Panstwowego-Zakladu-Higieny, 54(3), 2003, 263-267.
[6]. R. Zambiazi, and M.W. Zambiazi, Vegetable oil oxidation: effects of endogenous components, Revista da Sociedade Brasiliera de
Ciencia e Tecnologia de Alimentos, Camphinas, 34(1), 2000, 22-32.
[7]. D. Dzisiak, New oils reduced saturated and trans fats in processed foods, (Cereal Foods World 49(6), 2004, 331-333.
[8]. R. Przybylski ,B.E. McDonald, Develompent and Processing of vegetable oils for human nutrition in Illinios: The Oil Press
/AOCS,1995.
[9]. C. L. Lawton , H. J. Delargry , J. Brockman, R .C. Simith, and J.E. Blundell. The degree of saturation of fatty acids of fatty acids
influences in post ingestive satiety, British Journal of Nutrition, 83 (5), 2000,473-482.
[10]. F. Priego-Capote, J. Ruiz-Jiménez, J. Garcia-Olmo, and M. Luque de Castro, Fast method for the determination of total fat and trans
fatty-acids content in bakery products based on microwave-assisted Soxhlet extraction and medium infrared spectroscopy detection,
Analytica Chimica Acta, 517,2004,13-20.
[11]. F. Priego-Capote, J. Ruiz-Jiménez, and M. Luque de Castro, Identification and quantification of trans fatty acids in bakery products
by gas chromatography-mass spectrometry after focused microwave Soxhlet extraction, Food Chemistry, 100,2007,859-867.
[12]. W.W Christie, and X. Han, Lipid analysis: Isolation, separation, identification and lipidomic analysis, Oily Press Bridgewater UK,
2010.
[13]. B. Rozema, B. Mitchell, D. Winters, A. Kohn, D. Sullivan, and E. Meinholz, Proposed modification to AOAC 996.06, optimizing
the determination of trans fatty acids:presentation of data, Journal of AOAC International, 91,2008,92-97.
[14]. W.W. Christie, Preparation of ester derivatives of fatty acids for chromatographic analysis, Advanced in Lipid Methodology, 2,
1993, 69-111.
[15]. Brondz, Development of fatty acid analysis by high- performance liquid chromatography, gas chromatography and related
techniques, Analytica Chimica Acta, 465, 2002, 1-37.
[16]. P. Delmonte, and J.I. Rader, Evaluation of gas chromatographic methods for the determination of trans fat, Analytical Biochemistry,
389, 2007, 77-85.
[17]. M. Juárez, O. Polvillo, M. Contò, A. Fisco, S. Ballico, and S. Failla, Comparison of four extraction/methylation analytical methods
to measure fatty acid composition by gas chromatography in meat, Journal of Chromatography A, 1190, 2008, 327-332.
[18]. M. Careri, L. Elvin , A. Mangia, Liquid chromatography – UV determination and liquid chromatography – atmospheric pressure
chemical ionization mass spectrometric characterization of sitosterol and stigmasterol in soybean oil, Journal of Chromatography A,
935,2001, 249–257.
[19]. G. Contarini, M. Povolo, E. Bonfitto, S. Bererdi, Quantitative analysis of sterols in dairy products:experiences and remarks,
International Dairy Journal, 12, 2002,573–578.
[20]. T.A. Omar, and J. Salimon, Validation and application of a gas chromatographic method for determining fatty acids and trans fats in
some bakery products, Journal of Taibah University for Science, 7, 2013,56-63.
[21]. S. Hanwen, G. Xusheng, L. Yunkai, and W. Anbang, Application of accelerated solvent extraction in the analysis of organic
contaminants, bioactive and nutritional compounds in food and feed, Journal of Chromatography A, 1237, 2012, 1-23.
[22]. J. L Ezzell, B.E. Richter, W.D Felix., S. R. Black and J. E. Meikle, A comparison of Accelerated Solvent Extraction with
conventional solvent extraction for organophosphorus pesticides and herbicides, LC-GC, 13, 1995, 390 – 398.
[23]. J.M Snyder, J.W King. M. A. Jackson, Fat content for nutritional labeling by supercritical fluid extraction and on-line lipase
catalyzed reaction, J Chromatography A 750, 1996, 201-207.
[24]. S.L Аbidi, Chromatographic analysis of plant sterols in foods and vegetable oils, Journal of chromatography A, 935, 2001, 173-201
[25]. J.C. Miller. and J. N. Miller, Statistic for analytical chemistry, 3rd Ed., Ellis Horwood Ptr. Prentice Hall, 1993, 104 – 141.
[26]. L. R. Ellison (LGC, UK), A. Williams (UK), Quantifying Uncertainty in Analytical Measurement, Eurachem/CITAC Guide CG 4,
3rd Ed, 4-31, 2012.
[27]. K.W. Thomas, L.S. Sheldon, E.D. Pellizari, R.W. Handy, J.M. Roberts, and M.R. Berry, Testing duplicate diet sample collection
methods for measuring personal dietary exposures to chemical contaminants, J Expo Anal Environ Epidemio,l 7(1) 1997, 17-36.

C0532011019

More Related Content

What's hot (20)

Viewers also liked (17)

Similar to C0532011019 (20)

More from iosrphr_editor (20)

C0532011019