THERMAL DEGRADATION MONITORING OF BIODIESEL IN THE PRESENCE AND ABSENCE OF ALCOHOLIC EXTRACT OF JAMBOLAN (Syzygium cumini Lamarck) AND Cu²⁺ IONS BY ¹H NMR SPECTROSCOPY

Abstract

Biodiesel can be obtained from the transesterification reaction of triglycerides with short-chain alcohols in the presence of a catalyst. Some of the methyl esters present in biodiesel are unsaturated, which favors the oxidation process and can be slowed or suppressed with the addition of antioxidants. There are many options and varieties of synthetic antioxidants, such as tert butylhydroquinone (TBHQ) and butylhydroxytoluene (BHT). However, plant extracts, containing substances such as anthocyanins and flavonoids, with antioxidant properties can be used because they are effective in protecting biodiesel from oxidation. In this study, we evaluated the antioxidant behavior of jambolan pulp extract (Syzygium cumini Lamarck) in biodiesel, in the presence and absence of Cu²⁺, at different exposure times at a temperature of 110 °C with a flow rate of 10.00 L h^-1 of air using proton nuclear magnetic resonance (¹H NMR) spectroscopy. The high-intensity signals obtained by ¹H NMR spectroscopy showed that the biodiesel used was composed of 98.14% esters, with 35.35% saturated and 62.79% unsaturated esters. The low-intensity resonance signals showed that the main thermal degradation products were epoxides, aldehydes and hydroperoxides. The jambolan pulp extract reduced the oxidation reaction rate of biodiesel even in the presence of Cu²⁺ ions, demonstrating its potential use as an antioxidant.

Keywords:
biofuel; natural antioxidants; oxidative stability; reaction rate.

INTRODUCTION

Biodiesel production can be carried out through the transesterification reaction of a wide variety of raw materials, including vegetable oils, animal fats and waste frying oils with short-chain alcohol in the presence of a catalyst such as sodium or potassium hydroxide.¹ However, the fatty acid profile of these sources directly influences the physicochemical properties of the resulting biodiesel, especially its oxidative stability.

Triglycerides formed by unsaturated fatty acids, for example, confer greater susceptibility to oxidation, while saturated fatty acids tend to provide greater resistance to oxidative degradation. These characteristics are transferred to the esters of the biodiesel produced and chemical instability can be aggravated by environmental factors such as exposure to light, high temperatures, the presence of moisture and transition metal ions such as iron (Fe²⁺/Fe³⁺) and copper (Cu²⁺), which act as catalysts for the oxidation reactions of the biofuel esters. These reactions result in the formation of peroxides and secondary compounds that compromise fundamental physicochemical properties such as viscosity, acid value and other parameters, negatively affecting biodiesel performance in engines and its viability for medium and long-term storage.²,³

It is common to incorporate synthetic antioxidants, such as BHA (butylhydroxyanisole), BHT (butylhydroxytoluene) and TBHQ (tert-butylhydroquinone) to prolong the stability of biodiesel during transportation, storage and use, which act by inhibiting or delaying the oxidation reactions.⁴ These additives donate hydrogen atoms to the free radicals formed during the initiation stage of the oxidation reaction, preserving the essential physicochemical characteristics of the biofuel. However, in response to the growing demand for more sustainable and environmentally responsible solutions, the search for natural antioxidants obtained from renewable sources, although not yet commercially available for addition to biodiesel, such as the leaves, fruits and flowers of various plant species, has intensified. These sources are rich in bioactive compounds with antioxidant activity, including anthocyanins, carotenoids, and flavonoids.⁵

Wanasundara and Shahidi⁶ and Schaich⁷ describe that antioxidants can be grouped according to their mechanism of action: primary or secondary. Primary antioxidants can act by donating hydrogen to the free radical formed, restoring the biodiesel ester molecule and thereby delaying or inhibiting the initiation stage or interrupting the propagation stage of autoxidation due to the chemical nature of their molecules. Primary antioxidants include synthetic antioxidants such as BHA, BHT, and TBHQ and those obtained from plant sources, such as flavonoids. Secondary antioxidants, such as carotenoids, act as chelators for pro-oxidant or catalytic metal ions, inhibiting the catalysis of the oxidation reaction.⁶,⁸,⁹

In addition to their protective function against oxidative degradation, natural antioxidants offer additional advantages, such as reduced environmental impact, biodegradability and, in many cases, lower production costs, especially when using derivatives from agro-industrial waste. Therefore, the use of natural additives represents a promising strategy for stabilizing biodiesel, aligning technical performance with environmental and economic sustainability.¹⁰,¹¹

Jambolan (Syzygium cumini) is a fruit native to Indonesia, China and the Antilles, but this species has demonstrated excellent adaptation to Brazilian climate conditions.¹²,¹³ Brito et al.¹² identified a significant diversity of anthocyanins present in the jambolan peel, highlighting compounds such as delphinidin, cyanidin, petunidin, peonidin and malvidin. The antioxidant activity of the fruit is related to the presence of pigments, as well as vitamins, phenolic compounds, tannins and flavonoids. These constituents can minimize the concentration of free radicals formed during the oxidation stages, indicating the potential of jambolan extracts as natural antioxidants capable of slowing the oxidative degradation of biodiesel.¹³

Faria et al.¹³ determined the chromatographic and spectroscopic characteristics and composition of anthocyanins from jambolan fruit, obtained by high-performance liquid chromatography coupled with diode array detector and tandem mass spectrometry (HPLC DAD MS/MS), highlighting delphinidin 3,5-diglucoside, cyanidin 3,5-diglucoside, petunidin 3,5-diglucoside, peonidin 3,5-diglucoside among other compounds. In addition, they determined the carotenoid contents of cis-lutein, all-trans-zeaxanthin, 15-cis β carotene, 13-cis-β-carotene.

Branco et al.⁹ analyzed the phenolic compounds present in jambolan pulp and quantified the total levels of phenolic compounds and anthocyanins using ultraviolet-visible (UV-Vis) techniques. According to the authors, the quantification and chromatographic and spectroscopic characteristics of non-anthocyanin phenolic compounds in jambolan pulp obtained by HPLC-DAD revealed the presence of gallic acid, chlorogenic acid, rutin, myricetin and quercetin.

Branco et al.⁵ analyzed the alcoholic extract of jambolan fruit by high-performance liquid chromatography coupled with photodiode array detection (HPLC-PDA) and identified metabolites from the phenolic acid class, monomeric flavan-3-ols and their oligomers and polymers (proanthocyanidins), gallic acid, catechins and vanillic acid. Furthermore, analysis of the alcoholic extract of jambolan pulp by ultra-performance liquid chromatography-mass spectrometry (UPLC MS) revealed the presence of epicatechin/catechin, caffeoylquinic acid, chlorogenic acid, and anthocyanins such as cyanidin, malvidin, and peonidin. These compounds found in jambolan extract have antioxidant properties and, according to Branco et al.,⁵ can be applied to biodiesel to increase its oxidative stability, thereby extending the time between production and application in motor vehicles.

The objective of this study was to monitor the formation of oxidation products and the reaction rate during the thermal degradation of biodiesel, in the presence and absence of jambolan alcoholic extract and copper ions, using the Rancimat method and proton nuclear magnetic resonance (¹H NMR).

EXPERIMENTAL

Biodiesel production

The triglyceride transesterification reaction was carried out with 50.00% palm oil (S.S. Moratto Comércio de Insumos, São Paulo, SP, lot DE-0519-2023/28003) and 50.00% soybean oil (Cocamar^®, Maringá, PR, lot 22477), with absolute methanol (CH₃OH, FMaia, P.A. 99.80%, Belo Horizonte, MG) and potassium hydroxide (KOH, Cinética, 95.00%, Itapevi, SP) as the catalyst.

For every 100.00 g of triglycerides, 0.80 g of KOH dissolved in 50.00 mL of methanol was used. The mixture was heated to 60.0 °C under reflux and stirring for 2.00 h. The phases were separated in a separatory funnel and the biodiesel was subsequently washed in two stages, the first with a 1.00% m/m aqueous hydrochloric acid solution (LabSynth 36.50-38.00%, Diadema, SP) and then with water only. In the biodiesel washing process, the hydrochloric acid and water were at 80.0 °C and the procedure was carried out until neutral pH.

Anhydrous sodium sulfate (Anhydrol, 99.00%, Diadema, SP) was used to carry out the biodiesel drying process, which remained for 2.00 h in the oven at 140.0 °C. Subsequently, the sodium sulfate was cooled and mixed with stirring, remaining at rest for 1.00 h, followed by vacuum filtration, obtaining the biodiesel used for the analyses of the present research.

Natural jambolan pulp extract

The jambolan fruits (Syzygium cumini Lamarck) (National System for the Management of Genetic Heritage and Associated Traditional Knowledge (SisGen) A45BABD) were collected at the State University of Londrina at a point with geographic coordinates 23.328047 S 51.197556 W.

The jambolan pulp samples were oven-dried at 60.0 °C. They were then ground and vacuum-packed. An amount of 10.00 g of the dried samples were weighed and mixed in a beaker containing 250.00 mL of absolute ethyl alcohol (CH₃CH₂OH, FMaia, 99.80%, Belo Horizonte, MG) to prepare the extract. The beaker was then sealed with plastic film, wrapped in aluminum foil and stored in the dark for 48 h. The extract was then filtered using a glass funnel with quantitative filter paper and subsequently concentrated on a heating plate at approximately 60.0 °C until a final volume of less than 50.00 mL was reached. The final volume of the evaporated extract was transferred to a 50.00 mL volumetric flask, adjusted to the meniscus with absolute ethanol, wrapped in aluminum foil and stored in the refrigerator for later use.

Determination of phenolic compounds in extracts

The quantification of total phenolic compounds present in the alcoholic extract was performed by UV-Vis spectrometry using a Thermo Scientific instrument (model Evolution 60), at 760 nm, following the Folin-Ciocalteu method described by Branco et al.¹ A 2.0 M Folin-Ciocalteu phenol reagent (Sigma-Aldrich, Cotia, SP) and 7.50% m/v of the sodium carbonate solution (Anidrol, 99,5%, Diadema, SP) were used. The total phenol content was calculated and expressed in mg of gallic acid equivalent (GAE) per gram of dry matter.

Sample preparation

Biodiesel samples were prepared by adding 18.30 mL of jambolan pulp extract as recommended by Branco et al.,¹ corresponding to 42.80 mg of GAE to 100.00 g of biodiesel, all alcohol-free, stirring and letting it rest for 24 h. The alcohol from the extract was removed by evaporation using a hotplate at 60.00 °C.

Biodiesel samples containing Cu²⁺ ions were individually prepared by adding CuCl₂.2H₂O (Dinâmica, 99.00%, Jaraguá do Sul, SC) at a concentration of 9.96 × 10^-6 mol of cation to 40.00 g of biodiesel, with and without the addition of the natural extract. The samples containing Cu²⁺ ions were left to rest for 2.00 h before being subjected to the oxidative stability test.⁵

Rancimat method and electrical conductivity determination

Control biodiesel samples, and those containing the metallic ion, with and without natural extract, were subjected to heat treatment separately using a Metrohm Instruments Rancimat model 873 (Herisau, Switzerland). For each analysis, 7.50 g of the biodiesel sample was added to eight tests tubes, which were connected to the Rancimat instrument and subjected to controlled heating using two heating blocks set at 110.0 °C. Biodiesel oxidation was initiated by the continuous passage of atmospheric air at a constant flow rate of 10.00 L h^-1.

During the process, the volatile products formed by oxidative degradation were carried by the airflow to a measuring vessel containing ultrapure water (Milli-Q water) and a conductivity electrode. The volatile oxidation products transported by the air were collected in eight measuring vessels, from which the electrical conductivity was measured. The presence of volatile compound dissolved in water increases its electrical conductivity, a parameter monitored in real time to indicate the progress of the oxidation process of the biodiesel sample.

For each assay, one of the reaction vessels containing the biodiesel subjected to heating was removed according to the times shown in Table 1. For the last sample, in all cases, the time recorded was the moment when the conductivity of 200 μS was reached in the equipment. Each sample was then kept in a freezer at -5 °C for subsequent analysis by ¹H NMR spectroscopy.

Table 1
Sampling time for samples of biodiesel, biodiesel with jambolan extract, biodiesel with jambolan extract and Cu²⁺ ions, and biodiesel with Cu²⁺ ions

Sample	Sampling time / h
	1	2	3	4	5	6	7	8
Biodiesel	1.00	2.00	3.50	6.50	8.00	9.00	9.50	10.00
Biodiesel + jambolan	0.69	1.50	2.00	3.55	6.00	8.79	10.00	10.89
Biodiesel + jambolan + Cu2+	0.50	4.50	7.00	7.85	8.00	8.25	8.75	8.86
Biodiesel + Cu2+	0.95	1.69	2.64	3.18	-	-	-	-

Reaction rate (k)

The reaction rates (k) were determined by the adjusted natural logarithm (ln) of electrical conductivity versus time data provided by the Rancimat accelerated oxidative stability test at 110.0 °C, considering the first-order reaction (Equation 1):

(1) $$\ln \mathrm{\Lambda }=\ln {\mathrm{\Lambda }}_0-\mathrm{k}({\mathrm{t}}_{\mathrm{f}}-{\mathrm{t}}_{\mathrm{i}})$$

where Λ is the electrical conductivity at time t; Λ₀ is the initial conductivity, t_i and t_f correspond to the initial and final time, respectively.

The values of the reaction rates (k) were determined by the value of the slope of the fitted line (Equation 1).⁵

¹H NMR spectroscopy

¹H NMR spectroscopy analyses were performed to determine the proportion of esters in biodiesel and to identify the compounds present in samples with and without jambolan pulp extract and Cu²⁺, subjected to different exposure times to the oxidative process. It was used a high-resolution nuclear magnetic resonance spectrometer for spectral acquisition, operating at 9.4 T and 400 MHz (Bruker, Billerica, MA, USA). Samples were prepared by dissolving 50.00 μL of biodiesel in 600.00 μL of deuterated chloroform (CDCl₃, 99.80% with 0.05% (v/v) tetramethylsilane (TMS), Sigma-Aldrich, St. Louis, MO, USA). The parameters used were: single pulse, spectral width of 8012 Hz, 32 scans, relaxation delay of 1.00 s, pulse width of 90° and acquisition time of 4.089 s.

Determination of the proportion of esters in biodiesel by ¹H NMR spectroscopy

The proportions of unsaturated methyl esters (oleates, linoleates, linolenates) and saturated methyl esters present in biodiesel were obtained using the normalized areas of the signals present in the ¹H NMR spectra according to the equations proposed by Guillén and Ruiz.¹⁴ The total ester content (FAMEs) present in the biodiesel sample was determined according to the equation presented by Gelbard et al.¹⁵ and by Knothe,¹⁶ using the normalized areas of the methoxy protons (approximately 3.70 ppm) and the α-carbonyl methylene groups (approximately 2.30 ppm). The data from the ¹H NMR spectroscopy analyses, the determination of the proportion of esters in biodiesel and plot the graphs were performed using the OriginPro software, version 2025.¹⁷

Chromatographic analysis of biodiesel samples

Chromatographic analysis of the ester content was performed according to EN14103,¹⁸ using a Shimadzu GC2010 Plus chromatograph coupled to a QP2010 Ultra mass detector equipped with a Restek-RT2560 fused silica chromatographic column.

Statistical analysis

The statistical parameters of the models, including the coefficients of determination (R²) and correlation (r), applied in the adjustment of the data of the natural logarithm (ln) of electrical conductivity as a function of time and of the non-linear regression models, respectively, were determined using the Statistica software, version 13.4.0.33.¹⁹

RESULTS AND DISCUSSION

The obtained biodiesel samples were subjected to the oxidative stability test using the accelerated heating method via Rancimat. During the tests, 8 samples were collected at different time intervals, which were analyzed by ¹H NMR.

Thermal monitoring of biodiesel without additive

Figure 1 shows the ¹H NMR spectral signals of the control biodiesel sample (sample 0) and samples 2, 4, 6 and 8. On the left, the resonance spectra shows that most signals are located between 0 and 6.00 ppm, labeled from 1 to 9. The signals indicated by numbers 1 to 9 are those whose integrated areas were used to determine the saturated and unsaturated ester content in the biodiesel.

Figure 1
¹H NMR spectra with signals between 0.84-0.93 ppm (a); 0.93-1.02 ppm (b); 1.24-1.40 ppm (c); 1.56-1.68 ppm (d); 1.95-2.15 ppm (e); 2.26-2.34 ppm (f); 2.70-2.82 ppm (g); 3.64-3.70 ppm (h); 5.28-5.40 ppm (j) of the control biodiesel sample and the sample subjected to heating

On the right of the figure, magnifications of the signals are shown, separated by heating time. Since the intensity of the proton signal in the ¹H NMR spectra is directly related to their content in the sample, it is possible to monitor the evolution of the oxidation process through changes in signal intensities.

By integrating the areas of the 9 signals of the control sample (sample 0) and using the areas of the signals represented by the numbers 1, 2, 5, 6 and 7 and the equations proposed by Guillén and Ruiz,¹⁴ the proportions of the oleic, linoleic, linolenic and saturated acyl groups were determined. According to the same authors, the area of signal 1 corresponds to the methyl hydrogen atoms of saturated plus oleic and linoleic acyl groups; signal 2 to the methyl hydrogen atoms of the linolenic acyl groups; signal 5 to the methylene hydrogen atoms in the α position in relation to a double bond; signal 6 to the methylene hydrogen atoms in the α position, in relation to the carboxyl group; and signal 7 to the methylene hydrogen atoms in the α position in relation to two double bonds, also called bis-allylic protons.¹⁴

According to Gelbard et al.¹⁵ and Knothe,¹⁶ the yield of methyl esters was determined by the integration values of the protons of the methyl ester fraction, signal 8 (approximately 3.70 ppm) and of α-carbonyl methylene groups (at 2.30 ppm), corresponding to signal 6. The ester content of the biodiesel was estimated in an average total content of 98.14%. Furthermore, applying the equations proposed by Guillén and Ruiz¹⁴ and normalizing to the estimated total ester content, the results showed that the biodiesel obtained was composed of 35.35 ± 0.22% saturated methyl esters, 33.67 ± 0.26% methyl oleates, 25.54 ± 0.11% methyl linoleates and 3.58 ± 0.07% methyl linolenate esters, resulting in a total ester content of 98.14%. This value is consistent with that obtained by chromatographic analysis (99.31%), according to EN 14103.¹⁸ This composition complies with the current Brazilian standard, according to National Agency of Petroleum, Natural Gas and Biofuels (ANP) Resolution No. 920,²⁰ which requires a minimum ester content of 96.50%.

Figure 1a corresponds to signal 1, with a chemical shift between 0.85 and 0.91 ppm, which, according to Guillén and Ruiz,¹⁴ is produced by the overlap of triplet signals of protons from saturated acyl groups, ω-9, i.e., oleate, and ω-6, i.e., linoleate. The signal intensity is lower in the initial sample and higher in sample 8. This occurs because, according to Bacha et al.,²¹ polyunsaturated compounds are more susceptible to the loss of hydrogen from the C-H bond, leading to the formation of free radicals, increasing the rate of peroxidation. This interpretation is supported by Wagner et al.,²² who reported that susceptibility to peroxidation increases with the degree of unsaturation. The dissociation energy of the C-H bond ranges from 75.00 to 80.00 kcal mol^-1 at bis-allylic sites, whereas it is approximately 88.00 kcal mol^-1 at allylic positions and about 101.00 kcal mol^-1 in alkyl chains. According to Branco et al.,²³ it is reasonable to assume that the biodiesel oxidation reaction is primarily controlled by the initial hydrogen release from the C-H bond at the bis-allylic sites, since this process requires less energy than hydrogen abstraction from the allylic sites.

For saturated and monounsaturated esters, the oxidation reaction is slower because the hydrogens at the methyl and allylic sites require greater abstraction energy compared to those at the bis-allylic site. Consequently, when the propagation phase of the oxidation reaction begins, the observed increase in the proportion of saturated and monounsaturated esters is not due to their formation, but rather to the oxidative degradation of polyunsaturated methyl esters. This is particularly evident with the greater loss of methyl linoleates, which are initially present in higher concentrations than methyl linolenates. The area of signal 1 was used to determine the content of the compounds methyl oleate, methyl linoleate and methyl linolenates in the biodiesel sample.¹⁴

In Figure 1b, signals with chemical shifts ranging from 0.94 to 1.01 ppm, show a decrease in intensity from sample 0 to sample 8. Signal 2 corresponds to the methyl protons of the ω-3 acyl groups, and the difference in chemical shifts between the methyl proton signals is due to their proximity to the double bond of the carbon chain.¹⁴ The gradual decrease in the signal is due to the degradation of the monoand polyunsaturated compounds present in the biodiesel sample, which have allylic and bis-allylic sites that preferentially form free radicals, propagating the oxidation reaction. The area of signal 2 was used to determine the content of methyl oleate, linoleate and linolenates.

In Figure 1c, signal 3, with a chemical shift ranging from 1.23 to 1.33 ppm, increases in intensity with increasing exposure of the sample to heat treatment. According to Martínez-Yusta,²⁴ the signal in this range is related to acyl group protons.

Signal 4 (Figure 1d) corresponds to methylene protons in the β positions, relative to the carboxyl group, and appears between 1.58 and 1.65 ppm.¹⁴ The signal intensity increases with increasing heat exposure time.

Signal 5 (Figure 1e), between 1.95 and 2.10 ppm, is due to the presence of α-methylene protons in the sample, also called allylic protons.¹⁴ The area of signal 5 was used to determine the content of methyl oleate and methyl saturated compounds in the biodiesel sample.

Signal 6 (Figure 1f) corresponds to methylene protons in the α-position, relative to the carboxyl group and appears between 2.27 and 2.33 ppm, respectively.¹⁶ According to Morgenstern et al.,²⁵ the presence of esters produced by polyunsaturated fatty acids can be easily detected by the appearance of a signal in this range, belonging to the methylene group between two double bonds of a linoleic, linolenic or higher polyunsaturated fatty acid chain. Signal 6 had its area used to determine the methyl linoleate, oleate and saturates content¹⁴ and also in the determination of the total esters (FAMEs) content of biodiesel.¹⁵,¹⁶ It is interesting to note that these signals increase in intensity during the sample heating process due to an increase in the proportion of methyl saturates and oleates caused by the decrease in methyl linoleates and linolenates present in the sample.²³

Signal 7 (Figure 1g), which appears between 2.70 and 2.82 ppm, is due to the overlap of signals from methylene α-protons in relation to two double bonds, also called bis-allylic protons.¹⁴,²⁶ The presence of esters formed from polyunsaturated fatty acids can be easily detected by the appearance of a resonance at 2.72 ppm of the methylene group between two double bonds of a linoleic, linolenic or higher polyunsaturated fatty acid chain.²⁵ Signal 7 has its area used to determine the content of methyl linoleate and methyl oleate.¹⁴ Signal 8 (Figure 1h), between 3.65 and 3.68 ppm, is attributed to the protons on carbon atoms 1 and 3 of the glyceryl group.¹⁴ In this case, an increase in its intensity can also be observed with increasing heating time. According to Gelbard et al.¹⁵ and Knothe,¹⁶ its area can be used to estimate the FAME content of biodiesel.

The signal 9 (Figure 1i), between 5.31 and 5.40 ppm, is due to the olefinic protons of different acyl groups.¹⁴

Figure 2 shows the low-intensity signals of the ¹H NMR spectra, between 0 and 10.00 ppm of the control biodiesel sample (sample 0) and samples 2, 4, 6 and 8, subjected to heating times of 2.00; 6.50; 9.00 and 10.00 h respectively in the Rancimat apparatus.

Figure 2
¹H NMR spectra with low-intensity signals between 1.70-1.85 ppm (a); 2.40-2.60 ppm (b); 2.90-3.30 ppm (c); 3.92-4.04 ppm (d) 4.16-4.48 ppm (e); 5.45-5.60 ppm (f); 5.70-5.94 ppm (g); 5.95-6.15 ppm (h); 6.21-6.30 ppm (i); 6.56-6.68 ppm (j); 7.90-8.20 ppm (k) and 9.45-9.90 ppm (l) of the control biodiesel sample and samples subjected to heating at 110.0 °C, under aeration, for 10.00 h. Figures (a-1) represent enlarged views of the low-intensity signals of the five samples subjected to heating at different experimental times

The low-intensity signals, possibly from substances formed from the biodiesel oxidation reaction, were appropriately amplified. Since the intensity of the proton signal in the ¹H NMR spectra is directly related to its concentration in the sample, it is possible to monitor the evolution of the oxidation process through changes in signal intensities, highlighting the samples 6 and 8 with longer heating times. This may be attributed to the Rancimat test, which involves subjecting the sample to a temperature of 110 °C under continuous air flow, simulating accelerated oxidation conditions. During the Rancimat thermal degradation test, changes in the electrical conductivity of a receptor aqueous solution are monitored, indicating the formation of volatile compounds resulting from the oxidative decomposition of substances present in the biodiesel. However, an increase in the concentration of one or more esters in the sample, such as saturated and oleates methyl esters, can occur without their formation during the oxidation process. This happens due to the elimination, by the airflow to the electrical conductivity measurement cell, of polyunsaturated methyl esters that preferentially undergo decomposition, leading to a decrease in their concentration in the biodiesel samples.²³

In Figure 2a, signals can be observed between 1.67 and 1.85 ppm that can be attributed to acyl groups (-OCO-CH₂-CH₂-)²³ or even to alkenes and alkynes.²⁷ The 1.67 ppm signal decreases with heating time, suggesting the degradation of these compounds. In Figure 2b, signals between 2.43 and 2.49 ppm are observed for substances already present in sample 0, which increase in intensity with heating time. According to Martínez-Yusta et al.,²⁴ these signals can also be attributed to acyl groups. In the same figure, signals between 2.53 and 2.58 ppm are observed that were not present in sample 0 but were formed by thermal decomposition; their intensity increases in samples 6 and 8, which were subjected to 9.00 and 10.00 h of heating at 110.0 °C, respectively. According to Guillén and Ruiz,²⁸,²⁹ Martínez-Yusta et al.²⁴ and Mantovani et al.,³⁰ these low-intensity signals can be attributed to protons from monoand diepoxide structures. In Figure 2c, the signals between 2.85 and 3.30 ppm appear with greater intensity only in samples 6 and 8, which can also be attributed to epoxide groups. In Figure 2d, signals between 3.92 and 4.04 ppm are observed in sample 8, after 10.00 h of heat treatment, which, according to the same authors, can be attributed to epoxy structures. Among the epoxides, the following can be highlighted: 9,10-12,13-diepoxyoctadecanoate, 9,10-epoxy-octadecanoate, 9,10-epoxy-12-octadecenoate and 12,13-epoxy-9-octadecenoate.³¹,³²

In Figure 2e, signals between 4.12 and 4.23 ppm, present in the control sample, increase in intensity with the time of exposure to heat treatment and can be attributed to the presence of free glycerol, mono-, di-, and triacylglycerols,³³,³⁴ while signals between 4.24 and 4.40 ppm, not present in the control sample, can be attributed to the formation of methine proton (CH-OOH) from hydroperoxides.³⁵,³⁶

Chemical shift signals between 5.45 and 6.68 ppm (Figures 2f 2j), which are not present in the control sample, can, according to Martínez-Yusta et al.,²⁴ be attributed to compounds containing -CH=CH-CH=CH- structures, such as (Z,E)-with conjugated double bonds associated with hydroperoxides (OOH) or (E,E)-with conjugated double bonds associated with hydroperoxides (OOH). Furthermore, according to the same authors, the signal at 6.08 ppm (Figure 2h) may be related to the presence of a conjugated double bond belonging to a ketone group. Between 7.90 and 8.20 ppm (Figure 2k), there are no signals present in the control sample (0), which can be attributed to hydroperoxides formed from the reaction of unsaturated esters with oxygen during heat treatment.³⁰,³⁶

Finally, signals in the region of 9.45 to 9.90 ppm (Figure 2l) can be attributed to the presence of aldehydes such as (E,E)-2,4-alkadiene; 4,5-epoxy-2-alkene; 4-hydroxy-(E)-2-alkene; (Z,E)-2,4-alkadiene and n-alkanols, in agreement with literature data.²⁴,²⁸,²⁹,³⁷,³⁸

According to Lillard and Day³⁹ and Bacha et al.,²¹ the decomposition of hydroperoxides leads to the formation of alkoxy radicals (RO^•), which, in turn, can form a wide variety of compounds such as aldehydes, ketones, and alcohols. However, no signals were observed between 3.30 and 3.74 ppm that could correspond to the presence of primary alcohols, such as methanol (3.30 ppm) and ethanol (3.74 ppm), nor were any signals observed at 2.38 ppm indicating the presence of free fatty acids.²⁴,³³ The presence of oxidation products, such as epoxides, hydroperoxides and aldehydes, can affect the storage and use of biodiesel as a fuel in motor vehicles.²³

Jambolan extract

An alcoholic extract of jambolan pulp with antioxidant properties was produced and added to the biodiesel to slow down the biodiesel oxidation reaction. The extract was analyzed for total phenol content using the Folin-Ciocalteu method. The total phenol content, expressed in mg gallic acid equivalent (GAE), was 11.68 mg GAE g^-1 (dry mass), a value lower than that found by Branco et al.⁹

Eight samples of biodiesel with jambolan extract and biodiesel containing Cu²⁺ ions and extract were collected at different time intervals to monitor the degradation products formed during heat treatment by low-intensity ¹H NMR signals.

Thermal monitoring of low-intensity signals in biodiesel with jambolan extract

Figure 3 shows the ¹H NMR spectra, with adequately magnified low-intensity signals, of possible compounds formed during the oxidation reaction of biodiesel containing jambolan alcoholic extract, without heat treatment (sample 0) and of samples containing the extract subjected to 1.50 (sample 2); 3.55 (sample 4); 8.79 (sample 6) and 10.89 h (sample 8) of heating in the Rancimat apparatus. It is important to highlight that all the amplified signals, in all the bands presented in Figure 2, also appeared in the ¹H NMR spectra of the biodiesel sample containing the jambolan extract, although only some of them were highlighted in Figure 3.

Figure 3
¹H NMR spectra of low-intensity signals between 2.88-3.24 ppm (a); 3.90-4.10 ppm (b); 4.10-4.50 ppm (c); 5.87-5.95 ppm (d); 6.00-6.40 ppm (e) and 9.45-9.65 ppm (f) of the biodiesel sample with jambolan extract with and without heat treatment

The signals between 2.85 and 3.00 ppm that appear in Figure 3a, even in sample 0, also appeared in Figure 2c (samples 6 and 8), but now with greater intensity even in sample 0. This may indicate the incorporation of substances present in the extract that can produce signals in this range.⁴⁰ Signals between 3.05 and 3.20 ppm appear in samples 4, 6, and 8 subjected to 110.0 °C for 8.79 h (sample 6) and 10.89 h (sample 8). Similar behavior was observed in Figure 2c, where the sample did not contain the extract used.

According to Branco et al.⁹ and Faria et al.,¹³ jambolan extract is a source of phenolic compounds, with flavonoids being one of the most important classes, which provide ¹H NMR signals in the range of 3.90-4.00 ppm.⁴⁰ The signals observed between 3.90 and 3.97 ppm in Figure 3b, which appear even without thermal degradation, may be correlated with the incorporation of substances present in the jambolan extract.

The signals between 3.99 and 4.05 ppm, observed in samples subjected to greater thermal degradation (Figure 3b), according to Martínez-Yusta et al.²⁴ and Mantovani et al.,³⁰ can be attributed to epoxides structures. As in the samples presented in Figure 2, no signals indicative of the presence of primary alcohols (3.30 and 3.74 ppm) or free fatty acids (2.38 ppm) were observed in Figure 3.²⁴,³³

As discussed in Figure 2e, the signals present in Figure 3c between 4.10 and 4.21 ppm can be attributed to the presence of free glycerol, mono, di and triglycerides³³,³⁴ and, signals between 4.24 and 4.40 ppm, attributed to hydroperoxides that formed due to the heat treatment of the samples.³⁰,³⁶ Chemical shift signals between 5.67 and 6.35 ppm (Figures 3d and 3e) can be attributed to conjugated unsaturated substances associated with hydroperoxides and the signal at 6.08 ppm (Figure 3e) may be related to the presence of a conjugated double bond belonging to a ketone group.²⁴,²⁷ The signals observed in the region between 9.45 to 9.65 ppm in Figure 3f can be attributed to the presence of aldehydes originated from the radical oxidation reaction of biodiesel.²⁴,²⁷-²⁹,³⁷,³⁸

Biodiesel samples with and without jambolan extract and those containing extract and copper ions were prepared to evaluate the antioxidant effect of jambolan extract and its behavior in the presence of Cu²⁺ ions. These samples were also subjected to oxidative stability testing using the accelerated heating method (Rancimat). As the test of biodiesel containing Cu²⁺ ions lasted only 3.18 h, only four samples were collected.

Figure 4 shows the ¹H NMR spectra of appropriately magnified low-intensity signals, of possible compounds formed during the oxidation reaction of biodiesel containing Cu²⁺ ions and jambolan alcoholic extract, without heat treatment (sample 0) and of samples subjected to 4.50 (sample 2), 7.85 (sample 4), 8.25 (sample 6) and 8.86 h (sample 8) of heating in the Rancimat apparatus. All the amplified signals, in all the bands presented in Figure 2, also appeared in the ¹H NMR spectra of the biodiesel sample containing the jambolan extract and Cu²⁺ ions, but we highlight only a few of them in Figure 4.

Figure 4
¹H NMR spectra of low-intensity signals between 1.74-1.92 ppm (a); 3.10-3.16 ppm (b); 3.90-4.05 ppm (c); 4.14-4.41 ppm (d); 5.52-6.30 ppm (e); 6.03-6.30 ppm, (f); 6.51-6.60 ppm (g); 7.90-8.05 (h) and 9.50-9.80 ppm (i) of the biodiesel sample containing jambolan extract and Cu²⁺ ions with and without heat treatment

In the biodiesel sample with the addition of jambolan extract and Cu²⁺, signals between 1.68 and 1.90 ppm were observed (Figure 4a) with more intensity in the control biodiesel sample (sample 0). These signals are also present in the biodiesel sample without heat treatment (Figure 2a) and they are attributed to acyl groups.²⁸,²⁹ The signals observed in Figure 4b between 3.10 and 3.14 are of lower intensity than those observed in Figure 2c, indicating a lower formation of this type of degradation product. Due to the presence of carotenoids, such as β-carotene, lycopene and lutein, as well as anthocyanins in the jambolan extract,⁹ the extract also acts as a secondary antioxidant, that is, as a singlet oxygen quencher, delaying the oxidation reaction, even in the presence of copper ions.⁶,⁸ At the same time, flavonoids, which are multifunctional antioxidants, are also good metal ion chelators, inhibiting the initiation of oxidation by forming bonds with metal ions, reducing the redox potential and stabilizing the oxidized form of the metal ion.⁵,⁸,¹¹ This also explains why the biodiesel oxidation reaction in the presence of jambolan extract and Cu²⁺ lasted 8.86 h of thermal heating.

As in Figures 2 and 3, no signals indicative of primary alcohols (3.30 and 3.74 ppm) or free fatty acids (2.38 ppm) were observed in Figure 4.²⁴,²⁷,³³

Figure 4c, as in Figure 3b, shows a set of signals between 3.90 and 3.95 ppm that appear even without thermal degradation, but do not appear in the biodiesel samples (Figure 2d), which may be correlated with the presence of jambolan extract, which contains compounds that inhibit the catalytic action of Cu²⁺. As discussed in Figure 2e, the signals observed in Figure 4d, between 4.12 and 4.21 ppm, can be attributed to the presence of triacylglycerols, diacylglycerols and monoacylglycerols.²⁷,³⁴ Signals between 4.24 and 4.40 ppm, not observed in the control sample, can be attributed to the formation of hydroperoxides.²⁴,²⁷,³⁰,³³ There was an increase in the intensity of the signals observed in Figure 4e, referring to the range of 5.50-5.60 ppm, indicating that the presence of copper influenced the degradation even with the addition of the jambolan extract in terms of the formation of hydroperoxides, however more intense and in shorter exposure times than that observed in Figure 2f. The behavior in the range of 5.7 to 5.8 ppm was similar.

Signals between 5.01 and 6.10 ppm and between 6.20 and 6.30 ppm (Figure 4f), as well as signals between 6.52 and 6.60 ppm (Figure 4g), which are not observed in the control sample, show lower intensity than those observed in Figure 3e for substances associated with hydroperoxides.

Low-intensity signals in the 8.00-8.10 ppm range observed in biodiesel samples in the presence of extract and Cu²⁺ (Figure 4h) can be attributed to hydroperoxides formed from the reaction of unsaturated esters with oxygen during heat treatment.³⁰,³⁵,³⁶

The signals observed in the region between 9.45 to 9.80 ppm in Figure 4, as well as in Figures 2 and 3, can be attributed to the presence of aldehydes.²⁴,²⁸,³⁷,³⁸

Thermal monitoring of low-intensity signals in biodiesel with Cu²⁺

Figure 5 shows the ¹H NMR spectra of the biodiesel sample containing Cu²⁺ ions subjected to 0.95 h (sample 1); 1.69 h (sample 2); 2.64 h (sample 3), and 3.18 h (sample 4) of heating in the Rancimat apparatus. All the amplified signals, in all the bands presented in Figure 2, also appeared in the ¹H NMR spectra of the biodiesel sample (Figure 5), however, in this case, all the samples underwent heat treatment.

Figure 5
¹H NMR spectra of low-intensity signals between 1.68-1.86 ppm (a); 2.40-2.58 ppm (b); 2.94-3.12 ppm (c); 3.98-4.04 ppm (d); 5.80-5.92 ppm (e); 6.12-6.21 ppm (f); 6.51-6.63 ppm (g); 8.04-8.22 ppm (h); 9.50-9.90 ppm (i) of the biodiesel samples containing Cu²⁺ ions with and without heat treatment

In Figure 5a, the biodiesel sample in the presence of Cu²⁺ showed signals between 1.70 and 1.90 ppm of greater intensity than those observed in Figure 2a, indicating greater degradation of compounds attributed to acyl groups (-OCO-CH₂-CH₂-),²⁴ due to the presence of the copper ion, which acts as a catalyst for the biodiesel oxidation reaction.

The signals in Figure 5b, between 2.40 and 2.60 ppm, are of greater intensity than those in Figure 2b, indicating greater formation of compounds with acyl groups, due to the reaction catalysis promoted by the copper ion.²⁷ The signals between 2.50 and 2.60 ppm, which were not observed in sample 0 (Figure 2b), were formed by thermal decomposition. According to Guillén and Ruiz,²⁸,²⁹ Martínez Yusta et al.,²⁴ and Mantovani et al.,³⁰ these signals can be attributed to protons from monoand diepoxide structures. However, due to the presence of copper ions, these substances were formed even in the sample subjected to 1 h of heat treatment.

As in the control biodiesel sample (Figure 2d), Figure 5 shows signals between 2.90 and 3.15 ppm (Figure 5c) and 3.98 and 4.04 ppm (Figure 5d), which can be attributed to epoxide structures.³¹,³² As in the other figures, no signals indicative of the presence of primary alcohols (3.30 and 3.74 ppm) or free fatty acids (2.38 ppm) were observed in Figure 5.²⁴,³³

As already discussed in Figures 2f-2j, chemical shift signals between 5.79 and 6.63 ppm, observed in Figures 5e-5g, can be attributed to conjugated unsaturated substances associated with hydroperoxides (OOH),²⁴ with only variations in intensity.

The signals observed in the range of 8.00-8.10 ppm (Figure 5h), also observed in the biodiesel sample (Figure 2k), can be attributed to hydroperoxides that form from the reaction of unsaturated esters with oxygen during heat treatment.³⁰,³⁶

The signals observed in the region between 9.50 to 9.80 ppm (Figure 5i), as well as in Figures 2l, 3f and 4i, can be attributed to the presence of aldehydes.²⁴,²⁸,²⁹,³⁷,³⁸ It should be noted that due to the presence of copper ions, a catalyst for the oxidation reaction, the signals appeared with a shorter heating time and, in some bands, with greater intensity when compared to those shown in Figure 2.

Oxidation reaction rate (k)

Considering the formation of thermal degradation products, obtained via ¹H NMR, and the electrical conductivity data of the biodiesel samples, with and without jambolan extract, and those containing extract and Cu²⁺ ions, it was possible to monitor the behavior of the oxidation reaction rate (k).

The k values, in h^-1, were determined from the adjusted data of the natural logarithm (ln) of electrical conductivity vs. time, considering the first-order reaction, as described by Equation 1, with coefficients of determination (R²) ranging from 0.93 ≤ R² ≤ 0.99. These values can be considered acceptable for the oxidation reaction rate involving biodiesel, which has a complex chemical composition and also indicate that the choice of reaction kinetics first-order was adequate.¹¹

Biodiesel stability can be compromised by several physical and chemical factors, including hydrolysis, exposure to light radiation, thermal decomposition and contamination by impurities, especially catalytic metal ions. Fazal et al.⁴¹ demonstrated that biodiesel is more corrosive than diesel. This is crucial due to the transfer of metal ions to biodiesel, compromising its quality and oxidative stability.

Metallic contaminants can originate from copper heat exchangers used in biodiesel production, storage tanks or ferrous alloys present in materials during transportation, where contamination occurs through direct contact with the container surface or even through metallic sediments from the oxidation process.⁴² These agents promote chemical reactions that lead to the formation of undesirable compounds, such as peroxides, free fatty acids, aldehydes, ketones and polymers, which negatively affect the physicochemical properties and quality of the biofuel.¹¹

However, beyond these factors, biodiesel stability is directly related to the type of methyl esters it contains, especially the presence of unsaturation in the carbon chains. Unsaturated methyl esters, such as methyl oleate (monounsaturated), methyl linoleate (diunsaturated) and methyl linolenate (triunsaturated), exhibit varying degrees of susceptibility to oxidation. This vulnerability stems from the presence of hydrogen atoms in the allylic and, especially, bis-allylic positions, located between conjugated double bonds.³⁰

In these regions, hydrogen atoms are more easily abstracted, forming free radicals, initiating the formation of hydroperoxides and, subsequently, degradation products. Polyunsaturated esters are more reactive than monounsaturated esters. This trend is reflected in the increase in reactivity with the number of unsaturations, with methyl linolenate being the most susceptible due to the presence of three conjugated double bonds.³⁰

Figure 6 shows the behavior of the biodiesel oxidation reaction rate (k) in h^-1, as a function of time (h), in the presence and absence of jambolan extract (Figure 6a) and of biodiesel containing jambolan extract in the presence and absence of Cu²⁺ ions (Figure 6b). The points represent the experimental data, and the lines the data adjusted by the non-linear regression model, Y = a(b^x)(x^c), with the coefficients a, b and c adjusted using a convergence criterion of 1.00 × 10^-6, with 2 to 4 iterations, without weighting, with standard error ranging from 0.03 ≤ SE ≤ 0.31 and correlation coefficients between 0.99 ≤ r ≤ 1.00 indicating the good quality of the adjusted regression models.

Figure 6
Biodiesel oxidation reaction rate in the absence (′) and presence (π) of jambolan extract (a) and in the presence of Cu²⁺ with (*) and without (•) the extract (b). The points represent the experimental data, and the lines correspond to the non-linear regression model fit

It can be observed that the reaction rate declines due to the oxidation process of biodiesel subjected to the Rancimat test at 110.0 °C. This occurs because some of the unsaturated methyl esters present in the sample, which have bis-allylic bonds, preferentially undergo degradation, resulting in a decrease in k values due to the reduced content of these esters involved in the oxidation process. In the presence of jambolan extract, which has antioxidant properties, the effect is reversed because, at the beginning of the oxidation process, the free radicals formed are restored by the phenolic compounds present in the extract, slowing the oxidative process.

Figure 6b shows the behavior of the biodiesel oxidation reaction rate in the presence of Cu²⁺ ions with and without jambolan extract. The biodiesel oxidation reaction rate in the presence of Cu²⁺ ions decreases more rapidly because the degradation of the esters present is greater at the beginning of the reaction, undergoing a more pronounced decay, stabilizing after 3.50 h of testing. This occurs because, according to Spacino et al.,⁸ some transition metals, such as copper, interact with the unsaturations present in biodiesel esters through metal-olefin bonds. This explains why the Cu²⁺ ion exhibits a higher oxidation reaction rate (Figure 6b), as this ion has a high capacity to weaken the double bond of unsaturated methyl esters. Furthermore, in the presence of molecular oxygen, even at reduced pressures (pO₂), the reduced metal can act as a conductor, transferring an electron to oxygen to form the superoxide radical or its conjugate acid, the perhydroxyl radical.⁸ These species serve as a source of singlet oxygen, which, in turn, reacts with unsaturated esters to form hydroperoxides that form alkoxy and hydroxy radicals, rapidly propagating the ester oxidation reaction, forming a wide variety of compounds.³⁰,³⁵

Figure 6b shows the behavior of the biodiesel oxidation reaction rate in the presence of Cu²⁺ ions and jambolan extract, which contains phenolic compounds that act as primary antioxidants. The phenolic compounds donate hydrogen atoms, restoring biodiesel ester molecules, interrupting or inhibiting autoxidation and also acting as secondary antioxidants, chelating Cu²⁺ ions and inhibiting their action as a catalyst for the oxidation reaction.

CONCLUSIONS

The results showed that jambolan pulp extract can be used as an inhibitor of the biodiesel oxidation reaction, without significant changes in its antioxidant properties, while the presence of Cu²⁺ ions catalyzes the oxidation reaction.

To complement the data obtained, the reaction kinetics were evaluated, indicating that biodiesel samples containing jambolan pulp extract and Cu²⁺ ions showed lower oxidation reaction rates compared to those without the extract.

¹H NMR spectroscopy analysis proved to be a suitable method for determining the ester composition of biodiesel and for monitoring the oxidation reaction products at different sampling times. Biodiesel samples containing jambolan pulp extract and Cu²⁺ ions showed lower oxidation reaction rates compared to those without the extract.

ACKNOWLEDGMENTS

Laboratory of Chemometrics in Natural Sciences (UEL), Multiuser Research Laboratory Center - especially ESPEC (UEL), CNPq (303588/2021 7), and CAPES are acknowledged.

DATA AVAILABILITY STATEMENT

The authors state that all data are available in the text.

REFERENCES

• ¹ Branco, I. G.; Romagnoli, E. S.; Gonçales-Filho, J.; Mantovani, A. C. G.; Borsato D.; Rev. Cienc. Agron. 2025, 56, 1. [Crossref]
  » Crossref
• ² Rêgo, D. M. G.; da Silva, L. C.; Oliveira, M. N. M.; Melo, R. P. F.; Carmo, S. K. S.; Research, Society and Development 2020, 9, e-321997263. [Crossref]
  » Crossref
• ³ Cherwoo, L.; Gupta. I.; Flora, G.; Verma, R.; Kapil, M.; Arya, S. K.; Ravindran, B.; Khoo, K. S.; Bhatia, S. K.; Chang, S. W.; Nagamcharussrivichai, C.; Ashokkumar, V.; Sustainable Energy Technologies and Assessments 2023, 60, 103503. [Crossref]
  » Crossref
• ⁴ Kimura, M.; Savada, F. Y.; Romagnoli, E. S.; Chendynski, L. T.; Borsato, D.; Rev. Virtual Quim. 2019, 11, 1097. [Crossref]
  » Crossref
• ⁵ Branco, I. G.; Campos, J. W.; Sousa, E. G.; Silva, N. F.; Gonçales-Filho, J.; Pauli, E.; Romagnoli, E. S.; Mantovani, A. C. G.; Borsato, D.; Quim. Nova 2025, 48, e-20250099. [Crossref]
  » Crossref
• ⁶ Wanasundara, P. K. J. P. D.; Shahidi, F. In Bailey’s Industrial Oil and Fat Products; Shahidi, F., ed.; John Wiley & Sons Inc.: Hoboken, 2005, ch. 11. [Crossref]
  » Crossref
• ⁷ Schaich, K. M. In Bailey’s Industrial Oil and Fat Products; Shahidi, F., ed.; John Wiley & Sons Inc.: Hoboken, 2005, ch. 7. [Crossref]
  » Crossref
• ⁸ Spacino, K. R.; Angilelli, K. B.; Silva, H. C. In Mathematical Modeling of Biofuel: Synthesis and Storage; Angilelli, K. B.; Chendynski, L. T.; Mantovani, A. C. G., eds.; Springer: Cham, 2020, ch. 8.
• ⁹ Branco, I. G.; Moraes, I. C. F.; Argandonã, E. J. S; Madrona, G. S.; Santos, C.; Ruiz, A. L. T. G.; Carvalho, J. E.; Haminiuk, C. W. I.; Ind. Crops Prod. 2016, 89, 255. [Crossref]
  » Crossref
• ¹⁰ Chendynski, L. T.; Cordeiro, T.; Messias, G. B.; Mantovani, A. C. G.; Spacino, K. R.; Zeraik, M. L.; Borsato, D.; Fuel 2020, 261, 116379. [Crossref]
  » Crossref
• ¹¹ Clemente, M. A. J.; Branco, I. G.; Romagnoli, E. S.; Gonçales Filho, J.; Mantovani, A. C. G.; Chendynski, L. T.; Borsato, D.; Fuel 2023, 253, 129276. [Crossref]
  » Crossref
• ¹² de Brito, E. S.; de Araujo, M. C. P.; Alves, R. E.; Carkeet, C.; Clevidence, B. A.; Novotny, J. A.; J. Agric. Food Chem. 2007, 55, 9389. [Crossref]
  » Crossref
• ¹³ Faria, A. F.; Marques, M. C.; Mercadante, A. Z.; Food Chem. 2011, 126, 1571. [Crossref]
  » Crossref
• ¹⁴ Guillén, M. D.; Ruiz, A.; Eur. J. Lipid Sci. Technol. 2003, 105, 688. [Crossref]
  » Crossref
• ¹⁵ Gelbard, G.; Brès, O.; Vargas, R. M.; Vielfaure, I.; Schuchardt, U. F.; J. Am. Oil Chem. Soc. 1995, 72, 1239. [Crossref]
  » Crossref
• ¹⁶ Knothe, G.; J. Am. Oil Chem. Soc. 2000, 77, 489. [Crossref]
  » Crossref
• ¹⁷ OriginPro, version 2025; OriginLab Corporation, Northampton, USA, 2025.
• ¹⁸ EN14103: Determination of Total FAME and Linolenic Acid Methyl Ester in FAME with AC Biodiesel All in One Solution, European Committee for Standardization: Berlin, 2012.
• ¹⁹ Statistica, version 13.4.0; Statsoft Inc., Tulsa, USA, 2018.
• ²⁰ Agência Nacional do Petróleo, Gás Natural e Biocombustíveis (ANP); Resolução ANP No. 920, de 4 de abril de 2023; Estabelece a Especificação do Biodiesel e as Obrigações quanto ao Controle da Qualidade a Serem Atendidas pelos Agentes Econômicos que Comercializem o Produto em Território Nacional; Diário Oficial da União (DOU), Brasília, No. 66, de 05/04/2023, p. 51. [Link] accessed in March 2026
  » Link
• ²¹ Bacha, K.; Ben-Amara, A.; Vannier, A.; Alves-Fortunato, M.; Nardin, M.; Energy Fuels 2015, 29, 4345. [Crossref]
  » Crossref
• ²² Wagner, B. A.; Buettner, G. R.; Burns, C. P.; Biochemistry 1994, 33, 4449. [Crossref]
  » Crossref
• ²³ Branco, I. G.; Campos, J. W.; Silva, N. F.; Gonçales-Filho, J.; Romagnoli, E. S.; Mantovani, A. C. G.; Angilelli, K. B.; Borsato, D.; Biofuel 2026, 17, 104. [Crossref]
  » Crossref
• ²⁴ Martínez-Yusta, A.; Goicoechea, E.; Guillén, M. D.; Compr. Rev. Food Sci. Food Saf. 2014, 13, 838. [Crossref]
  » Crossref
• ²⁵ Morgenstern, M.; Cline, J.; Meyer, S.; Cataldo, S.; Energy Fuels 2006, 20, 1350. [Crossref]
  » Crossref
• ²⁶ Parker, T.; Limer, E.; Watson, A. D.; Defernez, C. M.; Williamson, C. D.; Kemsley, E. K.; TrAC, Trends Anal. Chem. 2014, 57, 147. [Crossref]
  » Crossref
• ²⁷ Pavia, D. L.; Lampman, G. M.; Kriz, G. S.; Introduction to Spectroscopy: A Guide for Students of Organic Chemistry, 3^rd ed.; Thomson Learning: Belmont, 2001.
• ²⁸ Guillén, M. D.; Ruiz, A.; Eur. J. Lipid Sci. Technol. 2005, 107, 36. [Crossref]
  » Crossref
• ²⁹ Guillén, M. D.; Ruiz, A.; J. Sci. Food Agric. 2005, 85, 2413. [Crossref]
  » Crossref
• ³⁰ Mantovani, A. C. G.; Romagnoli, E. S; Gonçales-Filho, J.; Sartor, R. A.; Branco, I. G.; Tupan, L. F. S.; Valezi, D. F.; Mincache, A. J.; Borsato, D.; Angilleli, K. B.; ACS Omega 2025, 10, 37305. [Crossref]
  » Crossref
• ³¹ Goicoechea, E.; Guillen, M. D.; J. Agric. Food Chem. 2010, 58, 6234. [Crossref]
  » Crossref
• ³² Le Gresley, A.; Ampem, G.; Grootveld, M.; Percival, B. C.; Naughton, D. P.; Food Funct. 2019, 10, 7952. [Crossref]
  » Crossref
• ³³ Ferreira, A. G. A.; Lima, W. X. P.; Rial, R. C.; Microchem. J. 2024, 207, 112226. [Crossref]
  » Crossref
• ³⁴ Anderson, L. A.; Franz, A. K.; Energy Fuels 2012, 26, 6404. [Crossref]
  » Crossref
• ³⁵ Mantovani, A. C. G.; Chendynski, L. T.; Galvan, D.; Macedo, F. C.; Borsato, D.; Di Mauro, E.; Fuel 2020, 274, 117833. [Crossref]
  » Crossref
• ³⁶ Alonso-Salces, R. M.; Holland, M. V.; Guillou, C.; Food Control 2011, 22, 2041. [Crossref]
  » Crossref
• ³⁷ Tyl, C. E.; Brecker, L.; Wagner, K.; Eur. J. Lipid Sci. Technol. 2008, 110, 141. [Crossref]
  » Crossref
• ³⁸ Guillén, M. D.; Ruiz, A.; Trends Food Sci. Technol. 2001, 12, 328. [Crossref]
  » Crossref
• ³⁹ Lillard, D. A.; Day, E. A.; J. Am. Oil Chem. Soc. 1964, 41, 549. [Crossref]
  » Crossref
• ⁴⁰ Cangeloni, L.; Bonechini, C.; Leone, G.; Consumi, M.; Andreassi, M.; Magnani, A.; Rossi, C.; Tamasi, G.; Foods 2022, 11, 2495. [Crossref]
  » Crossref
• ⁴¹ Fazal, M. A.; Jakeria, M. R.; Haseeb, A.; Rubaiee, S.; Energy 2017, 135, 220. [Crossref]
  » Crossref
• ⁴² Chendynski, L. T.; Romagnoli, E. R.; Mantovani, A. C. G.; Kimura, M.; Marquesan, L. C. M.; Borsato, D.; J. Braz. Chem. Soc. 2019, 30, 90. [Crossref]
  » Crossref

Edited by

• Associate Editor handled this article:
  Livia Cristina R. M. da Frota

Publication Dates

• Publication in this collection
  03 July 2026
• Date of issue
  2026

History

• Received
  11 Nov 2025
• Accepted
  16 Mar 2026
• Published
  24 Mar 2026