, Volume 82, Issue 1, pp 271–278 | Cite as

Direct Analysis of Synthetic Phenolic Antioxidants, and Fatty Acid Methyl Ester Stability in Biodiesel by Liquid Chromatography and High-Resolution Mass Spectrometry

  • Marcella Casagrande
  • Chadin Kulsing
  • Jalal T. Althakafy
  • Clarisse M. S. Piatnicki
  • Philip J. MarriottEmail author
Part of the following topical collections:
  1. 50th Anniversary Commemorative Issue


Methods to identify and quantify synthetic phenolic antioxidants, 3-tert-butyl-4-hydroxyanisole (BHA), butylated hydroxytoluene (BHT), tert-butyl-hydroquinone (TBHQ) and propyl gallate (PG), in biodiesel samples by using reversed-phase liquid chromatography (LC) were developed. Using a C18 phase with LC and UV detection showed co-elution between BHT and fatty acid methyl esters (FAME) in the biodiesel sample, whereas an alkyl phenyl modified stationary phase resulted in good separation of all antioxidants from the fatty acid matrix, and allowed more accurate quantification of antioxidants in biodiesel samples. The latter column was applied for further study. Calibration curves for the four antioxidants were constructed, and the limit of detection estimated. Good calibration linearity was observed over the investigated concentration range of 10–80 ppm, with correlation coefficients (R2) ranging from 0.9986 to 0.9995 for all antioxidants. LOD values of 0.010, 0.015, 0.0125 and 0.030 ppm, and recoveries of 70 ± 2, 85 ± 2, 103 ± 2 and 92 ± 4% for PG, TBHQ, BHA and BHT at injected concentrations of 35 ppm were established, respectively. The method was applied for quantification of antioxidants in biodiesel without addition of spiked antioxidants, then for biodiesel spiked with the four antioxidants, and a commercial source of biodiesel with BHT addition. Identification of FAME in the biodiesel samples was performed by using an instrument capable of ultra-high performance LC hyphenated with an electrospray Orbitrap mass spectrometer (UHPLC–ESI-OrbitrapMS). The stability of antioxidants and FAME in different samples was then investigated. Total FAME C18 content decreased to 52 ± 4% w/w after 1 week, and 29 ± 6% w/w after 8 weeks in the test sample without antioxidants; FAME content and antioxidant composition were stable in the samples with antioxidants added, even after 8 weeks exposure to sunlight.


Antioxidants Biodiesel Liquid chromatography Phenyl stationary phase Q-Exactive Quadrupole Orbitrap mass spectrometer 





Butylated hydroxytoluene


Electrospray ionisation


Liquid chromatography


Mass spectrometer


Propyl gallate


Synthetic phenolic antioxidants




High-performance liquid chromatography




Biodiesel is an alternative renewable fuel comprising long-chain mono-alkyl esters (fatty acid esters); it is commonly obtained by reaction between short-chain alcohols (such as methanol or ethanol) and vegetable oil/animal fat in presence of an acid or basic catalyst [1, 2]. This fuel presents some advantages compared to fossil fuels, e.g. lower toxicity, higher cetane number and flash point, higher lubricity and absence of sulfur and aromatic compounds. Furthermore, it can be used in diesel engines without the need of major modification [3]. Despite these advantages, biodiesel can be more easily oxidised or autoxidised under long-term storage than conventional diesel, with unsaturated more prone to oxidation than saturated esters [4], forming oxidation products, deposits or polymers that can damage vehicle engines and lead to their malfunction. Exposure to light, heat, humidity, oxygen and metals can accelerate this oxidation process; therefore, it is necessary to improve biodiesel stability as well as to monitor degradation processes [1, 2, 5]. In order to increase shelf life and delay biodiesel oxidation, synthetic phenolic antioxidants (SPAs) are often added to the biofuel, e.g. 3-tert-butyl-4-hydroxyanisole (BHA), butylated hydroxytoluene (BHT), tert-butyl-hydroquinone (TBHQ) and 3,4,5-trihydroxybenzoic acid propyl ester (propyl gallate, PG) [6, 7]. SPAs interrupt chain oxidation reactions by donating a proton, yielding stable free radicals which are unable to initiate or continue lipid oxidation. They must be added to biodiesel prior to use or storage, since they are not able to reverse the effects of oxidation processes which have already occurred [5, 8]. For conventional and alternatively derived middle distillate fuels, antioxidant ‘packages’ comprising single or multiple SPA are often added to improve the stability of biofuels such as those for aviation applications [8].

Biodiesel stability depends on storage conditions, which determines the extent of oxidation. The susceptibility of biodiesel to oxidation, when compared to fossil fuels, depends mainly on the content of unsaturated fatty acid esters that comes from the feedstock which is used to produce the biofuel. It also depends on the presence of natural antioxidants and storage conditions; if not properly stored, e.g. exposure to light, oxidation products such as aldehydes and alcohols may affect properties of the biodiesel. Thermal stability towards oxidative degradation relates the effect of temperature to formation of oxidation products, or reduction of biodiesel components as a result of oxidation; higher temperatures may enhance degradation rates. Storage stability may be described as the ability to resist change under long-term storage, maintaining the original physical and chemical characteristics. The standard method for oxidative stability analysis is the Rancimat method; nevertheless, many studies have been conducted in order to generate faster and more efficient methods for monitoring oxidation processes regarding storage time and sample aging [9, 10, 11, 12, 13].

A variety of analytical methods for biodiesel analysis have been reported, including electrochemical techniques [7, 14, 15, 16] and capillary electrophoresis [17, 18, 19], however gas chromatography (GC) [20, 21, 22, 23, 24] and high-performance liquid chromatography (HPLC) are the most widely used methods [6, 25, 26, 27, 28, 29, 30]. Analysis protocols for antioxidant composition of such samples often commence with sample extraction, often with large quantities of solvents, extended time, and an increased possibility of sample loss during extraction and/or derivatisation procedures. Fewer procedures report direct analysis of the unprocessed sample.

Methods based on GC offer good repeatability, high resolution and high peak capacity, and on-line coupling to MS permits reliable library searching, which can confirm detection of antioxidants in both biodiesel and edible oils without complex sample preparation methods [20, 21, 31, 32]. Application of comprehensive two-dimensional gas chromatography (GC × GC) further increases peak capacity, leading to enhanced resolution of antioxidants from the biodiesel matrix [33, 34, 35], as well as improved compound identification by increased quality of mass spectrometry matching, and potential for second dimension retention indices [36]. Derivatisation is often performed in order to improve peak quality (shape) and/or sensitivity in GC analysis [37, 38]. Alternatively, HPLC can be performed for antioxidant analysis, often without requiring derivatisation [6]. Fatty acid methyl ester (FAME) components in biodiesels can be adequately separated, e.g. under gradient elution in reversed-phase HPLC and FAME components may be identified by MS detection [39].

In this study, analysis of antioxidants in biodiesel was conducted by using HPLC–UV and HPLC–MS on the whole biodiesel sample. Limits of detection of antioxidants were calculated for the UV method. Antioxidants were identified by injecting authentic standards, and FAME identification was performed by using ultra-high performance liquid chromatography (UHPLC) but operated in conventional HPLC mode, hyphenated with a Q-Exactive Plus Orbitrap MS. Antioxidant and FAME composition in biodiesel samples, with and without antioxidant addition, were quantified after sample exposure to sunlight for different periods. Photo-oxidation (autoxidation is less rapid) and elevated temperature should be the primary oxidative processes arising from sunlight exposure [40].

Materials and Methods

Chemicals and Biodiesel Samples

A 37-component FAME mixture (47885-U) was obtained from Supelco (Sigma-Aldrich, St. Louis, MO). FAME with different chain length were abbreviated as FAME CX:Y, with the total carbon number and number of double bonds of X + 1 and Y, respectively. For example, FAME C18:0 and FAME C18:1 are methyl octadecanoate and methyl octadecenoate, respectively. GC-grade dichloromethane (DCM) was obtained from Merck Co. (Merck KGaA, Darmstadt, Germany). Water was distilled and deionised using a Milli-Q system (Millipore, Bedford, MA). Methanol (SupraSolv® grade) was obtained from Merck KGaA. Acetic acid (100% v/v) and phenol (used as internal standard) were purchased from Sigma-Aldrich and AnalaR (BDH Laboratory Supplies), respectively. The original FAME mixture (10,000 mg L−1 total concentration) was diluted to 1000 mg L−1 with DCM. tert-Butyl-hydroquinone (TBHQ) and butylated hydroxytoluene (BHT) were purchased from Fluka; 3-tert-butyl-4-hydroxyanisole (BHA) and propyl gallate (PG) were purchased from Sigma-Aldrich. Biodiesel samples (both pure and with BHT addition) were provided by ARFuels Co. Ltd. (Barnawartha, Australia). According to the company, the biodiesel sample has an added BHT concentration ranging from 600 to 800 ppm.

UV Degradation of Biodiesel

A total of 24 biodiesel samples (containing an initial concentration of 600–800 ppm BHT, quantified in this study to be 647 ± 10 ppm) were exposed to sunlight for different periods, ranging from 1 to 8 weeks. Three replicate samples were taken for analysis each week. The same series of experiments were also conducted on a corresponding set of 24 pure biodiesel samples (no added SPA), and 24 spiked with all 4 SPA; BHA, BHT, TBHQ and PG (700 ppm each). All samples were characterised by HPLC–UV. Samples were diluted 20 times in methanol, giving a nominal concentration of about 30–40 ppm in each antioxidant, prior to HPLC analysis.

Chromatographic Analysis and Instrumentation


Samples were analysed by using an Agilent 1220 Infinity LC system (Agilent Technologies, Mulgrave, Australia) with a reversed-phase C18 (Alltima 4.6 × 250 mm, 5 µm particle diameter; Agilent) or Zorbax SB-Phenyl column (StableBond Analytical 4.6 × 250 mm, 5 µm; Agilent). Although these columns have relatively large diameter particle size, they were found to be suited to the sample analysis reported, and so were used throughout this study. Samples were separated at 25 °C. 0.1% v/v acetic acid in water and methanol were used as mobile phases A and B, respectively. Gradient elution started at 50% v/v mobile phase B for 17 min and then linearly increased to 100% v/v B. The mobile phase content was held at 100% v/v B for 13 min then decreased to 50% v/v B. The column was equilibrated at 10% v/v B for 5 min resulting in the total time of 35 min prior to the next injection. Samples were diluted 20 times in methanol and injected (5 µL) at a flow rate of 0.6 mL/min for 35 min. Chosen wavelengths were 254, 260, 270, 280, 285, 290, 295 and 300 nm for selected chromatographic displays (detection bandwidth of 2 nm; reference off).

HPLC–Q-Exactive Orbitrap MS

Standards and samples were analysed by using a Dionex analytical ultra-high performance liquid chromatography (UHPLC) instrument, hyphenated with a Q-Exactive Plus Orbitrap mass spectrometer (ThermoFisher Scientific, Scoresby, Australia) which was equipped with a heated electrospray ionisation source (HESI), binary pump, autosampler and Quadrupole-Orbitrap. A Zorbax SB-Phenyl column (Agilent; as above) was employed as the stationary phase. To maintain the same retention and resolution obtained in the above HPLC–UV study, the same column was used, and the high pressures characteristic of UHPLC operation were not employed. The same temperature and mobile phase system as that used for HPLC–UV above were applied. The flow rate was 0.3 mL/min, and the injection volume was 15 µL. The HESI source was operated in both positive and negative modes. The source conditions were sheath gas 35, auxiliary gas 10, sweep gas 0, spray voltage 3.0 kV, capillary T 320 °C and auxiliary gas heater T 300 °C. The MS was operated in both positive and negative full scan modes (from 50 to 400 m/z), applied with MS parameters: resolution of 70,000 FWHM, AGC target of 1 × 106 and maximum injection time (IT) of 200 ms. The system was calibrated daily before analysis for both positive and negative modes.

Data Processing

Agilent OpenLAB CDS (ChemStation Edition) for LC and LC/MS systems and Microsoft Excel 2007 was used for data visualisation. Xcalibur 3.0.63 (ThermoFisher) software was used to control the Orbitrap instrument and for data analysis. TraceFinder 3.0 software (ThermoFisher) was used for identification, confirmation and quantification analysis with Microsoft Excel 2007.

Results and Discussion

Selection of Column and Wavelength with HPLC–UV

In order to investigate effects of column selection and wavelength, separation of biodiesel samples spiked with PG, TBHQ, BHA and BHT was performed. The detection wavelength of 280 nm was selected here to present an indication of the overall separation, since it resulted in an acceptable compromise signal for all the studied antioxidants and FAME. Although the C18 column resulted in good separation of SPA, co-elution between BHT and matrix interference components was observed; see Fig. 1a lower trace. Co-elution of SPA and matrix was still observed when aqueous/methanol mobile phase conditions were varied. Such co-elution leads to unreliable identification and quantification of BHT in practical samples. The alkyl-phenyl modified column was then tested, and resulted in good separation of all the antioxidants from the biodiesel matrix (refer to peaks 14 in Fig. 1a, upper trace). This column was selected for further studies. The presence of the phenyl group on the phase resulted in relatively stronger interaction between the phenyl stationary phase and the antioxidants, due to enhanced π–π interaction, compared to that of FAME. Hence the SPA moved to later retention, and toward the FAME matrix, but no overlap occurred. Variation of detection wavelength revealed that 270, 290, 290 and 280 nm gave the highest intensities for PG, TBHQ, BHA and BHT, respectively; see Fig. 1b for the corresponding peaks 14. Determination of limits of detection was performed according to these selected wavelengths for different antioxidants, using UV detection.

Fig. 1

a HPLC–UV analysis of four antioxidants (35 ppm each): PG (1), TBHQ (2), BHA (3) and BHT (4), Phenol (IS) in biodiesel matrix obtained at 280 nm by using alkylphenyl (above) and C18 (below) columns and b results obtained by using different wavelengths according to the same separation conditions used in a for the alkylphenyl column

Calibration Curves and Detection Limit

Calibration curves (concentration vs peak area ratio relative to the internal standard peak area at 280 nm) using HPLC–UV analysis were constructed for the four antioxidants by injecting the antioxidant mixture over the concentration range 10–80 ppm. The curves were linear over this range, with correlation coefficients between 0.9985 and 0.9994 for all four antioxidants (Fig. 2). This calibration range was selected to match the expected concentration range of antioxidants in real biodiesel samples after 20-fold dilution in methanol. The sensitivity of this method was sufficient, with limits of detection values estimated at 0.010, 0.015, 0.013 and 0.030 ppm for PG, TBHQ, BHA and BHT (observed at wavelengths of 270, 290, 290 and 280 nm, respectively), which have been identified from the lowest tested concentrations which resulted in a signal-to-noise ratio of the antioxidant peaks being ≥ 3. SPA concentrations considerably less than 10 ppm were tested for this calculation.

Fig. 2

Calibration curves generated in HPLC–UV analysis at 280 nm for PG, TBHQ, BHA and BHT (ad, respectively) response areas over the concentration range 10–80 ppm, vs the internal standard peak area. The corresponding calibration relationships are y = − 0.0992 + 0.2544x (R2 = 0.9995), y = 0.0432 + 0.0668 × (0.9986), y = − 0.0913 + 0.0674 × (0.9992) and y = − 0.0127 + 0.0373 × (0.9995), for a, b, c, d, respectively

Identification of Antioxidants in Biodiesel Samples

According to the calibration curves (Fig. 2), each antioxidant in spiked biodiesel samples was quantified. The SPA compounds were spiked into the biodiesel at a concentration of 700 ppm (after 20-fold dilution, 35 ppm was injected into the HPLC). The calculated concentrations for PG, TBHQ, BHA and BHT in the spiked biodiesel sample were 458 ± 11, 524 ± 14, 721 ± 17 and 556 ± 18 ppm, respectively. The error values were calculated standard deviations in the spiked biodiesel samples, calculated for three repeat sample preparations. Based on the injected spiked concentration of 35 ppm, recoveries of the four antioxidants in biodiesel sample matrix were 70 ± 2, 85 ± 2, 103 ± 2 and 92 ± 4%, respectively. The calculated concentration for BHT in the provided biodiesel sample to which BHT was added, was 647 ± 10 ppm, being within the range reported (600–800 ppm).

FAME and SPA Profiling by HPLC–Orbitrap MS

The four antioxidant peaks were detected using HPLC–Orbitrap MS with reduced background interference in the negative mode, as illustrated by the ion map plot of retention time vs m/z values (Fig. 3a). The four antioxidant peaks corresponded to the m/z values (212.20 g mol−1, 166.22 g mol−1, 180.25 g mol−1 and 220.36 g mol−1 for PG, TBHQ, BHA and BHT, respectively) of pseudo-molecular ions with loss of a proton (Table 1; see compounds 14). The mobile phase condition was the same as that applied in HPLC–UV analysis. However, the flow was 0.3 mL/min, compared with 0.6 mL/min used in HPLC–UV resulting in longer retention time (tR) observed in MS analysis. The slow flow facilitates spray formation in ESI–MS analysis. In addition to antioxidants, the FAME composition in biodiesel may be detected and identified. The negative ion mode gives many fewer FAME peaks displayed in the chromatogram (Fig. 3a), with many more components observed in positive ion mode (Fig. 3b). In the positive mode, according to the accurate mass values within ± 5 ppm mass accuracy, the protonated molecular ions were identified as shown in Table 1, with the ion map illustrated in Fig. 3b. Since increasingly hydrophobic compounds are more strongly retained on the hydrophobic phase, FAME with higher m/z values eluted later (located towards the top right corner of the ion map), and saturated FAME (more hydrophobic) eluted later than unsaturated FAME of similar molar mass (Table 1). These observed trends are indicative of a typical character of reversed-phase separation [30].

Fig. 3

HPLC–Q-Exactive Orbitrap MS ion map analysis of biodiesel spiked with PG, TBHQ, BHA and BHT (2 ppm each), labelled as peaks 14, respectively, in a negative and b positive modes with corresponding extracted ion chromatograms for the four antioxidants in c

Table 1

Analyte  analytical retention and MS data in biodiesel using HPLC–Q-Exactive Orbitrap MS


Retention time (min)

Exact mass [M + H]+ or [M − H]

Mass accuracy (ppm)

Possible compound




− 0.3





− 2.3





− 1.9








25.2, 27.6d


− 5.0

FAME C14:1


25.8, 27.8d


− 4.7

FAME C15:1


26.4, 28.6d


− 4.1

FAME C16:1


26.6, 28.2d


− 3.8

FAME C18:4


26.6, 28.5d


− 3.4

FAME C18:3


26.9, 29.0d


− 3.7

FAME C18:2


27.2, 29.5d


− 4.4

FAME C18:1




− 2.9

FAME C18:0




− 4.7

FAME C20:3


27.7, 30.0d


− 5.3

FAME C20:2


28.0, 30.6d


− 3.4

FAME C20:1


30.2, 31.5d


− 3.4

FAME C20:0




− 3.2

FAME C21:0

aCompound exact mass values calculated in negative ion mode

bCompound identity also confirmed by injection of authentic SPA compounds

cPseudo-molecular ion for the [M − H] ion

dCompound retention with >1 peak for an exact mass value indicates the presence of isomers

Biodiesel Stability Test

Stability of biodiesel components under exposure to sunlight was investigated, as a relatively simple exercise in forced oxidative degradation. The overlay chromatograms at different exposure periods of 0, 1, 4 and 8 weeks for biodiesel without antioxidant, biodiesel spiked with all four antioxidants, and the supplied biodiesel sample with BHT additive are shown in Fig. 4a–c. Without added antioxidant, the C18 FAME isomer content in biodiesel is significantly reduced to about 52 ± 4% w/w in the first week of exposure, decreasing to 29 ± 6% w/w after 8 weeks exposure (Fig. 5a). For samples containing antioxidants, the composition from 4 to 8 weeks did not vary significantly compared with the original assay at 0 weeks e.g. as illustrated for BHT (peak 4 in Fig. 4b, c with the corresponding content plotted in Fig. 5b) and other antioxidants (Fig. 5c). Some FAME with longer alkyl chains, such as FAME C18, slightly reduced to about 90% w/w of the original after 8 weeks exposure (Fig. 5a). Here, since the unsaturated C18 components are not resolved, and the saturated C18 is a minor component and elutes later, quantification is based on the unsaturated peak, which is more susceptible to oxidation. Although no particular temperature control was applied, the SPA were effective in reducing degradation; the effectiveness of SPA at elevated temperature is noted [4].

Fig. 4

a HPLC–UV analysis of biodiesel without antioxidant, b biodiesel spiked with PG, TBHQ, BHA and BHT (700 ppm of each SPA), and c the supplied biodiesel sample with BHT additive (600–800 ppm) after 0, 1, 4 and 8 weeks (black, green, purple and pink, respectively) under sunlight exposure. The labelled compounds are BHT (4) and FAME components (812) shown in Table 1 with the unknown oxidised species indicated in a

Fig. 5

Chemical profile in different biodiesel samples after exposure to sunlight up to 8 weeks: a total FAME C18 contents (compounds 812) in blank (times symbol), real biodiesel (open circle) and spiked biodiesel (open triangle) samples with and without BHT; b BHT (open square) content in real biodiesel sample and c PG (open diamond), TBHQ (open triangle), BHA (asterisk) and BHT (open square) contents in spiked biodiesel sample. The error bars were obtained from three replicates of the exposure experiments


This study develops techniques for direct analysis of antioxidants and FAME in biodiesel samples, to explore changes to FAME composition under exposure to sunlight as an accelerated oxidation condition. HPLC–UV was able to separate, detect and quantify four antioxidants without derivatisation with low LOD and good linearity of calibration curves. HPLC–Q-Exactive Orbitrap MS was applied for identification of antioxidants and FAME according to accurate mass analysis. Under exposure to sunlight as an accelerated oxidation process, the presence of antioxidant clearly reduced biodiesel degradation, where FAME content remained constant up to the maximum of the study period (8 weeks). The developed methods are expected to be applicable for reliable routine quantification of antioxidant in practical biodiesel samples, and investigation of biodiesel stability under different treatments. Compared to other methods the main advantage of the proposed methodology is the reduced time of analysis, because of the ease of sample preparation.



PJM acknowledges the Australian Research Council for a Discovery Outstanding Researcher Award; DP130100217. MC acknowledges CAPES Foundation for financial support. The authors acknowledge Agilent Technologies and ThermoFisher Scientific for provision of support for some of the facilities used in this study.

Compliance with Ethical Standards

Conflict of Interest

The authors declare that they have no conflict of interest.

Ethical Approval

This article does not contain any studies with human participants or animals performed by any of the authors.


  1. 1.
    Pullen J, Saeed K (2014) Factors affecting biodiesel engine performance and exhaust emissions—part I. Rev Energy 72:1–16CrossRefGoogle Scholar
  2. 2.
    Goulart LA, Teixeira ARL, Ramalho DA, Terezo AJ, Castilho M (2014) Development of an analytical method for the determination of tert-butylhydroquinone in soybean biodiesel. Fuel 115:126–131CrossRefGoogle Scholar
  3. 3.
    Ng J-H, Ng HK, Gan S (2010) Recent trends in policies, socioeconomy and future directions of the biodiesel industry. Clean Technol Environ Policy 12:213–238CrossRefGoogle Scholar
  4. 4.
    Dunn RO (2008) Antioxidants for improving storage stability of biodiesel. Biofuels Bioprod Biorefin 2:304–318CrossRefGoogle Scholar
  5. 5.
    Domingos AK, Saad EB, Vechiatto WWD, Wilhelm HM, Ramos LP (2007) The influence of BHA, BHT and TBHQ on the oxidation stability of soybean oil ethyl esters (biodiesel). J Braz Chem Soc 18:416–423CrossRefGoogle Scholar
  6. 6.
    Tagliabue S, Gasparoli A, della Bella L, Bondioli P (2004) Quali-quantitative determination of synthetic antioxidants in biodiesel. Riv Ital Sostanze Gr LXXXI:37–40Google Scholar
  7. 7.
    da Silva YP, Dalmoro V, Ruiz YPM, Capeletti LB, Mendonça CRB, dos Santos JHZ, Piatnicki CMS (2014) Biodiesel water in oil microemulsions: ferrocene as a hydrophobic probe for direct analysis by differential pulse voltammetry at a Pt ultramicroelectrode. Anal Methods 6:9212–9219CrossRefGoogle Scholar
  8. 8.
    Webster RL, Rawson PM, Evans DJ, Marriott PJ (2014) Synthetic phenolic antioxidants in conventional and alternatively-derived middle distillate fuels analysed by gas chromatography with triple quadrupole and quadrupole time of flight mass spectrometry. Energy Fuel 28:1097–1102CrossRefGoogle Scholar
  9. 9.
    Kivevele T, Huan Z (2015) Influence of metal contaminants and antioxidant additives on storage stability of biodiesel produced from non-edible oils of Eastern Africa origin (Croton megalocarpus and Moringa oleifera oils). Fuel 158:530–537CrossRefGoogle Scholar
  10. 10.
    Pereira GG, Alberici RM, Ferreira LL, Santos JM, Nascimento HL, Eberlin MN, Barrera-Arellano D (2015) A screening method to evaluate soybean oil-based biodiesel oxidative quality during its shelf life. J Am Oil Chem Soc 92:967–974CrossRefGoogle Scholar
  11. 11.
    Rashed MM, Kalam MA, Masjuki HH, Rashedul HK, Ashraful AM, Shancita I, Ruhul AM (2015) Stability of biodiesel, its improvement and the effect of antioxidant treated blends on engine performance and emission. RSC Adv 5:36240–36261CrossRefGoogle Scholar
  12. 12.
    Santos AGD, Souza LD, Caldeira VPS, Farias MF, Fernandes VJ Jr, Araujo AS (2014) Kinetic study and thermoxidative degradation of palm oil and biodiesel. Thermochim Acta 592:18–22CrossRefGoogle Scholar
  13. 13.
    Yang Z, Hollebone BP, Wang Z, Yang C, Brown C, Landriault M (2014) Storage stability of commercially available biodiesels and their blends under different storage conditions. Fuel 115:366–377CrossRefGoogle Scholar
  14. 14.
    Almeida JMS, Dornellas RM, Yotsumoto-Neto S, Ghisi M, Furtado JGC, Marques EP, Aucélio RQ, Marques ALB (2014) A simple electroanalytical procedure for the determination of calcium in biodiesel. Fuel 115:658–665CrossRefGoogle Scholar
  15. 15.
    Chýlková J, Tomášková M, Mikysek T, Šelešovská R, Jehlička J (2012) Voltammetric determination of BHT antioxidant at gold electrode in biodiesel. Electroanalysis 24:1374–1379CrossRefGoogle Scholar
  16. 16.
    Freitas HC, Almeida ES, Tormin TF, Richter EM, Munoz RAA (2015) Ultrasound-assisted digestion of biodiesel samples for determination of metals by stripping voltammetry. Anal Methods 7:7170–7176CrossRefGoogle Scholar
  17. 17.
    Spudeit DA, Piovezan M, Dolzan MD, Vistuba JP, Azevedo MS, Vitali L, Oliveira MAL, Costa ACO, Micke GA (2013) Simultaneous determination of free and total glycerol in biodiesel by capillary electrophoresis using multiple short-end injection. Electrophoresis 34:3333–3340CrossRefGoogle Scholar
  18. 18.
    Nogueira T, do Lago CL (2011) Determination of Ca, K, Mg, Na, sulfate, phosphate, formate, acetate, propionate, and glycerol in biodiesel by capillary electrophoresis with capacitively coupled contactless conductivity detection. Microchem J 99:267–272CrossRefGoogle Scholar
  19. 19.
    Piovezan M, Costa ACO, Jager AV, de Oliveira MAL, Micke GA (2010) Development of a fast capillary electrophoresis method to determine inorganic cations in biodiesel samples. Anal Chim Acta 673:200–205CrossRefGoogle Scholar
  20. 20.
    Okullo A, Ogwok P, Temu AK, Ntalikwa JW (2013) Gas chromatographic determination of glycerol and triglycerides in biodiesel from jatropha and castor vegetable oils. Adv Mater Res 824:436–443CrossRefGoogle Scholar
  21. 21.
    Hirschegger L, Schober S, Mittelbach M (2014) Efficient and sensitive method for the quantification of saturated monoacylglycerols in biodiesel by gas chromatography–mass spectrometry. Eur J Lipid Sci Technol 116:89–96CrossRefGoogle Scholar
  22. 22.
    Montpetit A, Tremblay AY (2016) A quantitative method of analysis for sterol glycosides in biodiesel and FAME using GC-FID. J Am Oil Chem Soc 93:479–487CrossRefGoogle Scholar
  23. 23.
    Bezerra KS, Antoniosi Filho NR (2014) Gas chromatographic analysis of free steroids in biodiesel. Fuel 130:149–153CrossRefGoogle Scholar
  24. 24.
    Farias AFF, Conceição MM, Cavalcanti EHS, Melo MAR, dos Santos IMG, de Souza AG (2016) Analysis of soybean biodiesel additive with different formulations of oils and fats. J Therm Anal Calorim 123:2121–2127CrossRefGoogle Scholar
  25. 25.
    Ahmed MA, Khan I, Hashima J, Musharraf SG (2015) Sensitive determination of glycerol by derivatization using a HPLC-DAD method in biodiesel samples. Anal Methods 7:7805–7810CrossRefGoogle Scholar
  26. 26.
    Fedosov SN, Fernandes NA, Firdaus MY (2014) Analysis of oil–biodiesel samples by high performance liquid chromatography using the normal phase column of new generation and the evaporative light scattering detector. J Chromatogr A 1326:56–62CrossRefGoogle Scholar
  27. 27.
    Haagenson DM, Perleberg JR, Wiesenborn DP (2014) Fractionation of canola biodiesel sediment for quantification of steryl glucosides with HPLC/ELSD. J Am Oil Chem Soc 91:497–502CrossRefGoogle Scholar
  28. 28.
    Allen SJ, Ott LS (2012) HPLC method for rapidly following biodiesel fuel transesterification reaction progress using a core-shell column. Anal Bioanal Chem 404:267–272CrossRefGoogle Scholar
  29. 29.
    Carvalho MS, Mendonça MA, Pinho DMM, Resck IS, Suarez PAZ (2012) Chromatographic analyses of fatty acid methyl esters by HPLC-UV and GC-FID. J Braz Chem Soc 23:763–769CrossRefGoogle Scholar
  30. 30.
    Brandão LFP, Braga JWB, Suarez PAZ (2012) Determination of vegetable oils and fats adulterants in diesel oil by high performance liquid chromatography and multivariate methods. J Chromatogr A 1225: 150–157CrossRefGoogle Scholar
  31. 31.
    Guo L, Xie M-Y, Yan A-P, Wan Y-Q, Wu Y-M (2006) Simultaneous determination of five synthetic antioxidants in edible vegetable oil by GC–MS. Anal Bioanal Chem 386:1881–1887CrossRefGoogle Scholar
  32. 32.
    Ding M, Zou J (2012) Rapid micropreparation procedure for the gas chromatographic-mass spectrometric determination of BHT, BHA and TBHQ in edible oils. Food Chem 131:1051–1055CrossRefGoogle Scholar
  33. 33.
    Marriott PJ, Chin S-T, Maikhunthod B, Schmarr H-G, Bieri S (2012) Multidimensional gas chromatography. TrAC Trends Anal Chem 34:1–21CrossRefGoogle Scholar
  34. 34.
    Mogollon NGS, Ribeiro FADL, Lopez MM, Hantao LW, Poppi RJ, Augusto F (2013) Quantitative analysis of biodiesel in blends of biodiesel and conventional diesel by comprehensive two-dimensional gas chromatography and multivariate curve resolution. Anal Chim Acta 796:130–136CrossRefGoogle Scholar
  35. 35.
    O’Neil GW, Culler AR, Williams JR, Burlow NP, Gilbert GJ, Carmichael CA, Nelson RK, Swarthout RF, Reddy CM (2015) Production of jet fuel range hydrocarbons as a coproduct of algal biodiesel by butenolysis of long-chain alkenones. Energy Fuel 29:922–930CrossRefGoogle Scholar
  36. 36.
    Jiang M, Kulsing C, Nolvachai Y, Marriott PJ (2015) Two-dimensional retention indices improve component identification in comprehensive two-dimensional gas chromatography of saffron. Anal Chem 87:5753–5761CrossRefGoogle Scholar
  37. 37.
    Nolvachai Y, Marriott PJ (2013) GC for flavonoids analysis: past, current and prospective trends. J Sep Sci 36:20–36CrossRefGoogle Scholar
  38. 38.
    Gao X, Williams SJ, Woodman OL, Marriott PJ (2010) Comprehensive two-dimensional gas chromatography, retention indices and time-of-flight mass spectra of flavonoids and chalcones. J Chromatogr A 1217:8317–8326CrossRefGoogle Scholar
  39. 39.
    Holčapek M, Jandera P, Fischer J, Prokeš B (1999) Analytical monitoring of the production of biodiesel by high-performance liquid chromatography with various detection methods. J Chromatogr A 858:13–31CrossRefGoogle Scholar
  40. 40.
    Knothe G (2007) Some aspects of biodiesel oxidative stability. Fuel Process Technol 88:669–677CrossRefGoogle Scholar

Copyright information

© Springer-Verlag GmbH Germany, part of Springer Nature 2018

Authors and Affiliations

  1. 1.Instituto de QuímicaUniversidade Federal do Rio Grande do SulPorto AlegreBrazil
  2. 2.Australian Centre for Research on Separation Science, School of ChemistryMonash UniversityMelbourneAustralia
  3. 3.Department of Chemistry, Faculty of Applied SciencesUmm Al-Qura UniversityMeccaSaudi Arabia
  4. 4.Department of Chemistry, Faculty of ScienceChulalongkorn UniversityBangkokThailand

Personalised recommendations