8

Lipid Oxidation: Measurement Methods

1. INTRODUCTION Dietary lipids, naturally occurring in raw food materials or added during food processing, play an important role in food nutrition and avor. Meanwhile, lipid oxidation is a major cause of food quality deterioration, and has been a challenge for manufacturers and food scientists alike. Lipids are susceptible to oxidative processes in the presence of catalytic systems such as light, heat, enzymes, metals, metalloproteins, and micro-organisms, giving rise to the development of off-avors and loss of essential amino acids, fat-soluble vitamins, and other bioactives. Lipids may undergo autoxidation, photo-oxidation, thermal oxidation, and enzymatic oxidation under different conditions, most of which involve some type of free radical or oxygen species (1, 2). Among these, only autoxidation and thermal oxidation are discussed here in detail. Autoxidation is the most common process leading to oxidative deterioration and is dened as the spontaneous reaction of atmospheric oxygen with lipids (3). The process can be accelerated at higher temperatures, such as those experienced during deep-fat frying, which is called thermal oxidation, with increases in free fatty acid and polar matter contents, foaming, color, and viscosity (4). Unsaturated fatty acids

Baileys Industrial Oil and Fat Products, Sixth Edition, Six Volume Set. Edited by Fereidoon Shahidi. Copyright # 2005 John Wiley & Sons, Inc.

357

358

LIPID OXIDATION: MEASUREMENT METHODS

are generally the reactants affected by such reactions, whether they are present as free fatty acids, triacylglycerols (as well as diacyglycerols or monoacylglycerols), or phospholipids (3). It has been accepted that both autoxidation and thermal oxidation of unsaturated fatty acids occurs via a free radical chain reaction that proceeds through three steps of initiation, propagation, and termination (5). A simplied scheme explaining the mechanism of autoxidation is given below:

Initiation:
initiator

LH

Propagation:
L LOO + O2 + LH LOO LOOH + L

Termination:
2 LOO LOO L + L + L Nonradical products

As oxidation normally proceeds very slowly at the initial stage, the time to reach a sudden increase in oxidation rate is referred to as the induction period (6). Lipid hydroperoxides have been identied as primary products of autoxidation; decomposition of hydroperoxides yields aldehydes, ketones, alcohols, hydrocarbons, volatile organic acids, and epoxy compounds, known as secondary oxidation products. These compounds, together with free radicals, constitute the bases for measurement of oxidative deterioration of food lipids. This chapter aims to explore current methods for measuring lipid oxidation in food lipids.

2. METHODS FOR MEASURING LIPID OXIDATION Numerous analytical methods are routinely used for measuring lipid oxidation in foods. However, there is no uniform and standard method for detecting all oxidative changes in all food systems (7). Therefore, it is necessary to select a proper and adequate method for a particular application. The available methods to monitor lipid oxidation in foods can be classied into ve groups based on what they measure: the absorption of oxygen, the loss of initial substrates, the formation of free radicals, and the formation of primary and secondary oxidation products (8). A number of physical and chemical tests, including instrumental analyses, have been employed in laboratories and the industry for measurement of various lipid oxidation parameters. These include the weight-gain and headspace oxygen uptake method for oxygen absorption; chromatographic analysis for changes in reactants;

MEASUREMENT OF OXYGEN ABSORPTION

359

iodometric titration, ferric ion complexes, and Fourier transform infrared (FTIR) method for peroxide value; spectrometry for conjugated dienes and trienes, 2-thiobarbituric acid (TBA) value, p-anisidine value (p-AnV), and carbonyl value; Rancimat and Oxidative Stability Instrument (OSI) method for oil stability index; and electron spin resonance (ESR) spectrometric assay for free-radical type and concentration. Other techniques based on different principles, such as differential scanning calorimetry (DSC) and nuclear magnetic resonance (NMR), have also been used for measuring lipid oxidation. In addition, sensory tests provide subjective or objective evaluation of oxidative deterioration, depending on certain details. 3. MEASUREMENT OF OXYGEN ABSORPTION 3.1. Weight Gain Consumption of oxygen during the initial stage of autoxidation results in an increase in the weight of fat or oil, which theoretically reects its oxidation level. Heating an oil and periodically testing for weight gain is one of the oldest methods for evaluating oxidative stability (9). This method requires simple equipment and directly indicates oxygen absorption through mass change. Oil samples are weighed and stored in an oven at a set temperature with no air circulation. To avoid the inuence of mass change by volatiles, samples can be preheated in an inert atmosphere. Samples are then taken out of the oven at different time intervals, cooled to ambient temperature, and reweighed; the weight gain is then recorded. The induction period can be obtained by plotting weight gain against storage time. In some cases, the time required to attain a 0.5% weight increase is taken as an index of oil stability (7, 9, 10). As a physical method for measuring lipid oxidation, the weight-gain method has several drawbacks such as discontinuous heating of the sample, which may give rise to non-reproducible results, and requiring long analysis time and intensive human participation (7). Nevertheless, this method offers advantages such as low instrumentation cost as well as a high capacity and processing speed of samples without limitation (7). Antolovich et al. (9) suggested that this technique may be extended to more sophisticated continuous monitoring of mass and energy changes as in thermogravimetry (TG)/differential scanning calorimetry (DSC). The weight-gain method can also be used for measuring antioxidant activity by comparing the results in the presence and absence of an antioxidant. Nevertheless, this method is useful only when highly unsaturated oils, such as marine oils and vegetable oils containing a high content of polyunsaturated fatty acids, are examined. 3.2. Headspace Oxygen Uptake In addition to the weight-gain method, oxygen consumption can be measured directly by monitoring the drop of oxygen pressure. Using headspace oxygen method, an oil sample is placed in a closed vessel also containing certain amount of oxygen at elevated temperatures, commonly around 100 C. The pressure reduction in

360

LIPID OXIDATION: MEASUREMENT METHODS

the vessel, which is due to the oxygen consumption, is monitored continuously and recorded automatically. The induction period as the point of maximum change in rate of oxygen uptake can be calculated (11). A commercial instrument for this method, known as Oxidograph, is available. In the Oxidograph, the pressure change in the reaction vessel is measured electronically by means of pressure transducers (7, 12). Oxygen consumption can also be measured by electrochemical detection of changes in oxygen concentration. However, the analysis of the graphical data obtained has been the bottleneck for this technique. The use of a semiautomatic polarographic method has been proposed as an improvement for evaluation of lipid oxidation by determination of oxygen consumption (13). As described by Genot et al. (13), this method is based on use of two oxygen meters with microcathode oxygen electrodes, coupled to a computerized data collection and processing unit. The headspace oxygen method is simple and reproducible and may be the best analytical method to evaluate the oxidative stability of fats and oils (14). Its application in measurement of lipid oxidation in food products other than fats and oils, however, is limited because protein oxidation also absorbs oxygen (15).

4. MEASUREMENT OF REACTANT CHANGE Lipid oxidation can also be assessed by quantitatively measuring the loss of initial substrates. In foods containing fats or oils, unsaturated fatty acids are the main reactants whose composition changes signicantly during oxidation. Changes in fatty acid composition provide an indirect measure of the extent of lipid oxidation (15). In this method, lipids are extracted from food, if necessary, and subsequently converted into derivatives suitable for chromatographic analysis (7). Fatty acid methyl esters (FAME) are the derivatives frequently used for determination of fatty acid composition, usually by gas chromatography (GC) (16). Similarly, iodine value, which reects the loss of unsaturation, can also be used as an index of lipid oxidation (17). Measurement of changes in fatty acid composition is useful for identication of lipid class and fatty acids that are involved in oxidation reactions (7). However, because the distribution of unsaturated fatty acids varies in different food systems, for instance, the highly unsaturated fatty acids being located predominantly in phospholipids of muscle foods, separation of lipids into neutral, glycolipid, phospholipid, and other classes may be necessary (7, 15). Moreover, it is an insensitive way of assessing oxidative deterioration. For comparison through calculation, oxidation of 0.4% polyunsaturated fatty acids to monohydroperoxides would represent a change of 16 meq oxygen/kg oil in peroxide value, whereas a change of less than 1.0 meq oxygen/kg oil could readily be detected by measuring peroxide value (12). Additionally, the application of this method is limited because of its inability to serve as an indicator of oxidation of more saturated lipids (7). Nevertheless, its usefulness for measuring oxidation of highly unsaturated oils cannot be underestimated.

MEASUREMENT OF PRIMARY PRODUCTS OF OXIDATION

361

5. MEASUREMENT OF PRIMARY PRODUCTS OF OXIDATION 5.1. Peroxide Value (PV) Lipid oxidation involves the continuous formation of hydroperoxides as primary oxidation products that may break down to a variety of nonvolatile and volatile secondary products (8, 15). The formation rate of hydroperoxides outweighs their rate of decomposition during the initial stage of oxidation, and this becomes reversed at later stages. Therefore, the peroxide value (PV) is an indicator of the initial stages of oxidative change (18). However, one can assess whether a lipid is in the growth or decay portion of the hydroperoxide concentration by monitoring the amount of hydroperoxides as a function of time (7). Analytical methods for measuring hydroperoxides in fats and oils can be classied as those determining the total amount of hydroperoxides and those based on chromatographic techniques giving detailed information on the structure and the amount of specic hydroperoxides present in a certain oil sample (8). The PV represents the total hydroperoxide content and is one of the most common quality indicators of fats and oils during production and storage (9, 18). A number of methods have been developed for determination of PV, among which the iodometric titration, ferric ion complex measurement spectrophotometry, and infrared spectroscopy are most frequently used (19).
5.1.1. Iodometric Titration Method Iodometric titration assay, which is based

on the oxidation of the iodide ion (I) by hydroperoxides (ROOH), is the basis of current standard methods for determination of PV (9). In this method, a saturated solution of potassium iodide is added to oil samples to react with hydroperoxides. The liberated iodine (I2) is then titrated with a standardized solution of sodium thiosulfate and starch as an endpoint indicator (7, 9, 20). The PV is obtained by calculation and reported as milliequivalents of oxygen per kilogram of sample (meq/kg). The ofcial determination is described by IUPAC (21). Chemical reactions involved are given below: ROOH 2H 2KI ! I2 ROH H2 O 2K I2 2NaS2 O3 ! Na2 S2 O6 2NaI Although iodometric titration is the most common method for measurement of PV, it suffers from several disadvantages. The procedure is time-consuming and laborintensive (18). As described by Ruiz et al. (18), the assay includes six steps: accurate weighing of the sample, dissolution of lipids in chloroform, acidication with acetic acid, addition of potassium iodide, incubation for exactly 5 minutes, and titration with sodium thiosulfate. This technique requires a large amount of sample and generates a signicant amount of waste (18, 22, 23). Furthermore, possible absorption of iodine across unsaturated bonds and oxidation of iodide by dissolved oxygen are among potential drawbacks of this method (7, 9). Besides, lack of sensitivity, possible interferences, and difculties in determining the titration endpoint

362

LIPID OXIDATION: MEASUREMENT METHODS

are also the main limitations (8, 23). To overcome these drawbacks, novel methods based on the same reaction have been developed, in which some other techniques are adopted as modication of the classical iodometric assay. Techniques such as colorimetric determination at 560 nm (24), potentiometric endpoint determination (25), and spectrophotometric determination of the I chromophore at 290 nm or 3 360 nm (26, 27) have been proposed. In addition, an electrochemical technique has been used as an alternative to the titration step in order to increase the sensitivity for determination of low PV by reduction of the released iodine at a platinum electrode maintained at a constant potential (7).
5.1.2. Ferric Ion Complexes Other chemical methods based on the oxidation of

ferrous ion (Fe2) to ferric ion (Fe3) in an acidic medium and the formation of iron complexes have also been widely accepted. These methods spectrophotometrically measure the ability of lipid hydroperoxides to oxidize ferrous ions to ferric ions, which are complexed by either thiocyanate or xylenol orange (23, 28, 29). Ferric thiocyanate is a red-violet complex that shows strong absorption at 500 510 nm (8). The method of determining PV by coloremetric detection of ferric thiocyanate is simple, reproducible, and more sensitive than the standard iodometric assay, and has been used to measure lipid oxidation in milk products, fats, oils, and liposomes (8, 23). The ferrous oxidation of xylenol orange (FOX) assay uses dye xylenol orange to form a blue-purple complex with a maximum absorption at 550600 nm (8). This method is rapid, inexpensive, and not sensitive to ambient oxygen or light (30). It can consistently quantify lower hydroperoxide levels; and good agreement exists between the FOX assay and the iodometric method (30). The FOX method has been successfully adapted to a variety of applications. However, because many factors, such as the amount of sample, solvent used, and source of xylenol orange, may affect the absorption coefcient, knowledge of the nature of hydroperoxides present in the sample, and careful control of the conditions used are required for accurate measurements (8).
5.1.3. Fourier Transform Infrared Spectroscopy (FTIR)

It has been recognized that hydroperoxides can quantitatively be determined by IR spectroscopy via measurement of their characteristic O-H stretching absorption band (31). An absorption band at about 2.93 mm indicates the generation of hydroperoxides, whereas the replacement of a hydrogen atom on a double bond or polymerization accounts for the disappearance of a band at 3.20 mm. The formation of aldehydes, ketones, or acids gives rise to an extra band at 5.72 mm. Furthermore, cis-, transisomerization and formation of conjugated dienes can be detected through the changes in the absorption band in the range of 10 mm to 11 mm (7). A rapid Fourier transform infrared spectroscopy (FTIR) method based on the stoichiometric reaction of triphenylphosphine (TPP) with hydroperoxides has been developed and successfully applied to determination of PV of edible oils (32). The hydroperoxides present in oil samples react stoichiometrically with TPP to produce triphenylphosphine oxide (TPPO), which has an intense absorption

MEASUREMENT OF PRIMARY PRODUCTS OF OXIDATION

363

band at 542 cm1 in the mid-IR spectrum (8, 18). The band intensity is measured and converted to peroxide value. The chemical reaction involved is given below: ROOH TPP TPP O ROH ! By using tert-butyl hydroperoxide spiked oil standards and evaluation of the band formed at 542 cm1, a linear calibration graph covering the range of 1100 PV was obtained (18). More recently, disposable polymer IR (PIR) cards have been used as sample holders where unsaturated oil samples oxidize at a fairly rapid rate (33). In the FTIR/PIR card method, warm air continuously ows over the sample allowing oxidation to be monitored at moderate temperatures. At periodic intervals, individual cards are removed and the FTIR spectra scanned (33). Another new FTIR approach uses ow injection analysis (FIA), which offers exact and highly reproducible timing of sample manipulation and reaction as well as a closed environment with oxygen and light being easily excluded (18). The FTIR spectroscopy is a simple, rapid, and highly precise method. It shows excellent correlation with the iodometric method and avoids the solvent and reagent disposal problems associated with the standard wet chemical method (18, 32). The FTIR method provides an automated, efcient and low-cost means of evaluating oxidation in oils undergoing thermal stress and has gained considerable interest for quality control in the industry (8, 20, 34). However, there is a need to characterize the spectral changes, assign wavelengths to more common molecular species produced, and access potential spectral cross interferences (20). Recently, an improved Fourier transform infrared attenuated total reectance (FTR-ATR) method using the whole FTIR spectral data instead of particular wavenumbers has been proposed (34). In addition to the three major methods discussed above, other techniques have also been employed in determination of PV, such as chemiluminescence and chromatography. Chemiluminescence method is based on detecting the chemiluminescent products generated during the reaction of hydroperoxides with substances such as luminol and dichlorouorescein (7, 35). This method was reviewd by Jimenez et al. (36). High correlations have been found between chemiluminescence and other standard methods, indicating that chemuliminescence could serve as an accurate tool for determination of PV (37). However, this method has low sensitivity to tert-butyl hydroperoxide, tert-butyl perbenzoate, diacyl peroxides, and dialkyl peroxides (35). Chromatographic techniques, mainly gas chromatography (GC) and high-performance liquid chromatography (HPLC), have also been employed for evaluation of lipid oxidation. These methods provide information about specic hydroperoxides, whereas other assays measure their total amount. Chromatographic methods require small amounts of sample, and interference from minor compounds other than hydroperoxides can be easily excluded (8). HPLC shows advantages over GC and has become a popular technique for hydroperoxide analysis. It operates at room temperature, thus decreases the risk of artifact formation, and no prior derivatization is required (8). A wide range of hydroperoxides can be analyzed using either normal or reverse-phase HPLC. Thus, hydroperoxides, the primary products

364

LIPID OXIDATION: MEASUREMENT METHODS

and intermediates in lipid oxidation reaction, provide an important parameter for evaluation of oxidation level. In addition, the inhibition of formation or action of these unstable species by antioxidants can be used as a means of assessing antioxidant activity (9). Measurement of hydroperoxides is also carried out in accelerated tests to establish the oxidative stability of a given oil. A case in point is the active oxygen method (AOM), in which air is bubbled through fat or oil held at 98100 C and PV is determined periodically (7, 38). The time required to reach a PV of 100 meq/kg is the AOM stability of the oil sample (7). This method is now considered outdated and is replaced by other standard methods in the industry, although product specications still routinely give AOM values (38). 5.2. Conjugated Dienes and Trienes It was discovered in 1933 that the formation of conjugated dienes in fats or oils gives rise to an absorption peak at 230235 nm in the ultraviolet (UV) region. In the 1960s, monitoring diene conjugation emerged as a useful technique for the study of lipid oxidation (9). During the formation of hydroperoxides from unsaturated fatty acids conjugated dienes are typically produced, due to the rearrangement of the double bonds. The resulting conjugated dienes exhibit an intense absorption at 234 nm; similarly conjugated trienes absorb at 268 nm (7). An increase in UV absorption theoretically reects the formation of primary oxidation products in fats and oils. Good correlations between conjugated dienes and peroxide value have been found (39, 40). Ultraviolet detection of conjugated dienes is simple, fast, and requires no chemical reagents and only small amounts of samples are needed. However, this

hydroperoxydiene

oxodiene

O O H

O Reduction hydroxydiene

O H conjugated triene and conjugated tetraene

Figure 1. Chemical reaction steps in conjugable oxidation products (COP) assay.

TABLE 1. Summary of Methods for Analysis of Primary Oxidation Products. Method Iodometric titration (PV) Ferric ion complexes (PV) Principle Reduction of ROOH with KI and measurement of I2 Reduction of ROOH with Fe2 and formation of Fe3 complexes Measurement Titration with Na2S2O3 Absorption at 500510 nm of the red complex with SCN Absorption at 560 nm of the blue-purple complex with xylenol orange Absorption at 542 cm1 of TPPO Chemiluminescence emission of oxidized luminol ROH derivatives Sensitivity %0.5-meq/kg fat %0.1-meq/kg fat Applications Fats and oils Fats, oils and food lipids

%0.5-meq/kg sample

All samples

FTIR (PV) Chemiluminescence (PV)

Reduction of ROOH with TPP Reaction with luminol in the presence of heme catalyst Reduction of ROOH to ROH and quantitation of ROH derivatives Estimation of conjugated dienes and trienes

%0.2-meq/kg fat %1 pmol

Fats and oils Fats and oils

GC-MS (PV)

UV spectrometry (conjugated dienes and trienes)

Absorption at 230234 nm and 268 nm

From ng to fg depending on technical details, amount of sample and detection system %0.2 meq/kg lipid

All samples

All samples

NOTE: The oxygen absorption measurement and loss of double bonds for fatty acid analysis are not considered as primary changes in this table. Adapted from (8).

366

LIPID OXIDATION: MEASUREMENT METHODS

method has less specicity and sensitivity than PV measurement (9, 12). Furthermore, the result may be affected by the presence of compounds absorbing in the same region, such as carotenoids (7). To avoid these interferences, an alternative spectroscopic method measuring conjugable oxidation products (COPs) has been proposed. In this method, hydroperoxides and some decomposition products are converted to more conjugated chromophores by reduction and subsequent dehydration (Figure 1). The concentrations of the resultant conjugated trienes and tetraenes are determined from their respective absorption at 268 nm and 301 nm and expressed as COP values (7, 12). Table 1 summarizes different methods available for analysis of primary oxidation products. Both chemical and instrumental methods are included in this table.

6. MEASUREMENT OF SECONDARY PRODUCTS OF OXIDATION The primary oxidation products (hydroperoxides) are unstable and susceptible to decomposistion. A complex mixture of volatile, nonvolatile, and polymeric secondary oxidation products is formed through decomposition reactions, providing various indices of lipid oxidation (5). Secondary oxidation products include aldehydes, ketones, alcohols, hydrocarbons, volatile organic acids, and epoxy compounds, among others. Methods for assessing lipid oxidation based on their formation are discussed in this section. 6.1. Thiobarbituric Acid (TBA) Test The thiobarbituric acid (TBA) test was proposed over 40 years ago and is now one of the most extensively used methods to detect oxidative deterioration of fat-containing foods (41). During lipid oxidation, malonaldehyde (MA), a minor component of fatty acids with 3 or more double bonds, is formed as a result of the degradation of polyunsaturated fatty acids. It is usually used as an indicator of the lipid oxidation process, both for the early appearance as oxidation occurs and for the sensitivity of the analytical method (42). In this assay, the MA is reacted with thiobarbituric acid (TBA) to form a pink MA-TBA complex that is measured spectrophotometrically at its absorption maximum at 530535 nm (Figure 2) (9, 43, 44). The extent of oxidation is reported as the TBA value and is expressed as milligrams
OH N HS N TBA OH MA TBA-MA adduct
Figure 2. Reaction of 2-thiobarbituric acid (TBA) and malonaldehyde (MA).

O O + H C O CH2 C H HN S N OH O

OH NH N H S

MEASUREMENT OF SECONDARY PRODUCTS OF OXIDATION

367

of MA equivalents per kilogram sample or as micromoles of MA equivalents per gram of sample. It must, however, be noted that alkenals and alkadienals also react with the TBA reagent and produce a pink color. Thus, the term thiobarbituric acid reactive substances (TBARS) is now used instead of MA. The TBA test can be performed by various procedures, among which four major types have frequently been employed. These include test on the whole sample, test on an aqueous or acid extract of sample, test on a steam distillate, and test on extracted lipid from a sample (45). The test on a steam distillate (distillation method) is the most commonly used method for determining TBA value. Tarladgis et al. (46) found that the distillation of an acidied sample was essential to liberate MA from precursor or bound forms, to produce maximal color development and, especially, to separate TBARS from the food matrix (44). Although the distillation method is the most popular TBA method, it is generally considered less accurate and reproducible than the method using food extracts (15). However, trends obtained in comparative studies always provide useful information that correspond with other measurements. Comparison of different TBA test procedures has been made by Hoyland et al. (46), Shahidi et al. (47), Pikul et al. (48), and Wang et al. (49). The TBA test is used frequently to assess the oxidative state of a variety of food systems, despite its limitations, such as lack of specicity and sensitivity (44). As already noted, many other substances may react with the TBA reagent and contribute to absorption, causing an overestimation of the intensity of color complex (44). Interferences may come from additional absorption of other alkanals, 2-alkenals, 2,4-alkdienals, ketones, ketosteroids, acids, esters, proteins, sucrose, urea, pyridines, and pyrimidines, also referred to as TBARS (43, 50). For instance, the reaction of TBA with various aldehydes leads to the development of a yellow chromogen (aldehyde-TBA adduct) with an absorption maximum at 450 nm, which overlaps with the pink peak at 532 nm resulting in erroneously high TBA values in certain cases (43, 45, 51). Furthermore, the presence of barbituric acid impurities in the TBA reagent may produce TBA-MA-barbituric acid and MA-barbituric acid adducts that absorb at 513 nm and 490 nm, respectively, indicating that thiobarbituric acid should be puried before use (43). In addition, nitrite can interfere in the TBA test, whereas sulfanilamide could be added to samples to avoid the interference when residual nitrite is present (52). In order to improve the specicity and sensitivity of the TBA test, several modications to the original TBA procedures have been proposed, including reduction of the heating temperature to stabilize the yellow color aldehyde-TBA complex (53), addition of antioxidants to sample in an attempt to prevent oxidation during the test (54), extraction of the MA prior to the formation of the chromogen (43), direct FTIR analysis of TBARS, and use of HPLC to separate the complex before measurement or to characterize the individual species of TBARS (9, 43). Despite it limitations, the TBA test provides an excellent means for evaluating lipid oxidation in foods, especially on a comparative basis. However, its use in bulk oils is less common than the so-called para-anisidine value (p-AnV) detailed below.

368

LIPID OXIDATION: MEASUREMENT METHODS

6.2. p -Anisidine Value (p -AnV) The p-anisidine value (p-AnV) method measures the content of aldehydes (principally 2-alkenals and 2,4-alkadienals) generated during the decomposition of hydroperoxides. It is based on the color reaction of p-methoxyaniline (anisidine) and the aldehydic compounds (55). The reaction of p-anisidine reagent with aldehydes under acidic conditions affords yellowish products that absorb at 350 nm (Figure 3) (7, 12). The color is quantied and converted to p-AnV. The p-AnV is dened as the absorbance of a solution resulting from the reaction of 1 g of fat in isooctane solution (100 ml) with p-anisidine (0.25% in glacial acetic acid) (12). This test is more sensitive to unsaturated aldehydes than to saturated aldehydes because the colored products from unsaturated aldehydes absorb more strongly at this wavelength (12). However, it correlates well with the amount of total volatile substances (55). The p-AnV is a reliable indicator of oxidative rancidity in fats and oils and fatty foods (56). A highly signicant correlation between p-AnV and avor scores and PV has been found (57). Nevertheless, some authors have indicated that p-AnV is comparable only within the same oil type because initial AnV varies among oil sources (58). For instance, oils with high levels of polyunsaturated fatty acids might have higher AnV even when fresh (59). This method is used less frequently in North America, but is widely employed in Europe (38), particularly as a part of the Totox number, as explained below. Caution
O H C C H Malonaldehyde (enolic form) OH C H + CH3O p-Methoxyaniline (p-anisidine) NH2

N CH3O

OH

NH2 CH3O

N CH3O

NH OCH3

Figure 3. Possible reactions between p-anisidine reagent and malonaldehyde.

MEASUREMENT OF SECONDARY PRODUCTS OF OXIDATION

369

must be exercised when performing this test because of toxicity of the anisidine reagent (55). 6.3. Totox Value The Totox value is a measure of the total oxidation, including primary and secondary oxidation products. It is a combination of PV and p-AnV: Totox value 2 PV p-AnV During lipid oxidation, it is often observed that PV rst rises, then falls as hydroperoxides decompose (38). PV and p-AnV reect the oxidation level at early and later stages of oxidation reaction, respectively. Totox value measures both hydroperoxides and their beakdown products, and provides a better estimation of the progressive oxidative deterioration of fats and oils (38). However, Totox value has no scientic basis because it is a combination of two indicators with different dimensions (7). Recently, Wanasundara and Shahidi used TBA values and dened TotoxTBA as 2PV TBA using the TBA test in place of the p-AnV assay (60). 6.4. Carbonyls The carbonyl compounds, including aldehydes and ketones, are the secondary oxidation products generated from degradation of hydroperoxides, and are suggested to be the major contributors to off-avors associated with the rancidity of many food products (9). The analysis of total carbonyl compounds, which is based on the absorbance of the carbonyl derivatives, provides another approach to measure the extent of lipid oxidation in fats and oils. In this method, the total carbonyl content is measured by a colorimetric 2,4-dinitrophenylhydrazone procedure. The carbonyl compounds formed during lipid oxidation are reacted with 2,4-dinitrophenylhydrazine (DNPH) followed by the reaction of the resulting hydrazones with alkali (Figure 4). The nal colored products are then analyzed spectrophotometrically
R R C H O + H2N N NO2
OH H2O

R NO2

H NO2 NO2

R C N N

R R C N N NO2

NO2

Figure 4. Reactions between carbonyls and 2,4-dinitrophenylhydrazine.

370

LIPID OXIDATION: MEASUREMENT METHODS

at a given wavelength (7, 15). Many variations of this method using an alternative solvent, reagent, wavelength, or workup have been reported. The determination of total content of carbonyls has been used in different oxidative stability studies. However, it has been criticized because the determination conditions cause degradation of hydroperoxides into carbonyl derivatives, giving erroneous results (58). Carbonyls produced from protein oxidation may also give rise to higher values than those expected from lipid oxidation alone. The addition of triphenylphosphine (TPP) prior to carbonyl determination has been proposed to avoid the interference from hydroperoxides. Hydroperoxides are reduced by TPP, and neither TPP nor TPPO, the oxidation products of TPP, interfere with the measurement of carbonyl content (61). In quality assessment of used frying fats, where short-chain carbonyls are already removed by distillation at the high temperature of the deep-frying, selectivity can be improved by determination of higher carbonyl compounds instead of the total carbonyls. HPLC is used to separate the DNPH derivatives of higher carbonyls from those of short-chain carbonyl compounds (62). Apart from detection of total carbonyl content, the analysis of individual carbonyl compounds has gained popularity for following lipid oxidation. Hexanal, one of the major secondary products formed during the oxidation of linoleic and other o6 fatty acids, serves as a reliable indicator of lipid oxidatin in foods rich in o6 fatty acids (7). A strong linear relationship was reported between hexanal content, sensory scores, and TBA values (63). Moreover, measurement of hexanal offers the advantage of analyzing a single, well-dened end product for antioxidant efciency studies (9). Hexanal can be quantied by chromatography (64) or as the intensity of the carbonyl band by NIR spectroscopy (65). Nevertheless, these methods may require volatilization of hexanal, whereas hexanal volatilization may be hindered due to covalent or other types of binding between hexanal and proteins in foods and, thus, may affect accurate hexanal quantications (66). More recently, an indirect enzyme-linked immunosorbanct assay (ELISA) has been developed for monitoring lipid oxidation through quantication of hexanal-protein adducts, which are recognized by polyclonal or monoclonal antibodies (66). Other carbonyl compounds, including propanal, pentanal, decadienal, etc., are also used for evaluating lipid oxidation in foods. For instance, propanal is a recommended indicator for lipid oxidation in foods that are high in o3 fatty acids, such as marine oils (67, 68). In general, it is essential to use appropriate indicators when assessing the oxidative deterioration of different food systems. 6.5. Oil Stability Index (OSI) During lipid oxidation, volatile organic acids, mainly formic acid and acetic acid, are produced as secondary volatile oxidation products at high temperatures, simultaneously with hydroperoxides (20, 69). In addition, other secondary products, including alcohols and carbonyl compounds, can be further oxidized to carboxylic acids (20). The oil stability index (OSI) method measures the formation of volatile acids by monitoring the change in electrical conductivity when efuent from oxidizing oils is passed through water (12). The OSI value is dened as the point of maximal change of the rate of oxidation, attributed to the increase of conductivity

MEASUREMENT OF SECONDARY PRODUCTS OF OXIDATION

371

by the formation of volatile organic acids during lipid oxidation (70). However, this method requires a somewhat higher level of oxidation (PV > 100) to obtain measurable results than other methods in which hydroperoxides are the most important products formed and detected (71). Therefore, to determine oil stability in the laboratory, especially for some oils that are stable under normal conditions, the oxidation process is accelerated by exposing oil samples to elevated temperatures in the presence of an excess amount of air or oxygen (72, 73). The OSI method differs from ambient storage conditions by using a ow of air and high temperatures to accelerate oxidation (71). The OSI is an automated development of the activeoxygen method (AOM), because both employ the principle of accelerated oxidation. Nevertheless, the OSI test measures the changes in conductivity caused by ionic volatile acids, whereas PV is determined in the AOM (7). Two pieces of commercially available equipment, the Rancimat (Metrohm Ltd.) and the Oxidative Stability Instrument (Omnion Inc.), are employed for determining the OSI value. Rancimat is a rapid automated method, which agrees well with the AOM (71). In the Rancimat assay, a ow of air is bubbled through a heated oil, usually at 100 C or above. For marine oils, temperatures as low as 80 C are often used. Volatile compounds formed during accelerated oxidation are collected in distilled water, increasing the water conductivity. The change of conductivity is plotted automatically and the induction period of the oil or the time taken to reach a xed level of conductivity is recorded (20, 74). The Rancimat assay enables continuous monitoring of the oxidation process. As reported by Farooq et al. (75), analysis by the Rancimat method is four to ve times more rapid than that by the AOM. Excellent correlation between Rancimat and conjugated dienes has been found (72). However, the main shortcoming of this method is that only eight samples can be included in each batch. Another appatatus, the Oxidative Stability Instrument, operates on the same principle as the Rancimat, and has the capacity of simultaneously analyzing up to 24 samples (20). Various modications have been proposed for assessing lipid oxidation by the OSI method. These include the use of auxiliary energies, such as microwaves to shorten the analysis time (72) and a combination of the OSI method with chromatography to obtain specic information about volatile products (76). The volatiles trapped during measurement by the Rancimat assay can be analyzed by headspace-GC (HS-GC) with FID and GC-MS for quantication of individual volatiles, thus improving the specicity of the assessment (76). Although the OSI method is useful for quality control of oils, it is not recommended for measurement of antioxidant activity for certain reasons. The high temperatures used do not allow reliable predictions of antioxidant effectiveness at lower temperatures. Volatile antioxidants may be swept out of the oil by the air ow under test conditions, and also the oils are severely deteriorated when endpoint is reached (12). 6.6. Hydrocarbons and Fluorescence Assay Formation of saturated hydrocarbons, especially short-chain (C1-C5) hydrocarbons such as ethane, propane, and pentane, can be measured for monitoring lipid oxidation when aldehydes are either absent or undetectable (7, 15). Pentane content,

372

LIPID OXIDATION: MEASUREMENT METHODS

O H C CH CHOH

RNH2

RN CH CH CHOH Amine-Malonaldehyde Adduct (Non-fluorescent)

O RN CH CH CH NHR Conjugated Schiff Base (Fluorescent)

H C CH2 C H Malonaldehyde

Figure 5. Reaction of lipid oxidation products such as malonaldehyde and amines.

determined by GC techniques, has been a useful parameter to assess rancidity of fats and oils as well as freeze-dried muscle foods (7, 15). Signicant correlations existed between pentane levels and rancid odor scores (15). It has been observed that the content of secondary oxidation products, such as malonaldehyde (MA), decreases with increased lipid oxidation, which can be explained by further reaction of MA with proteins. MA reacts with compounds containing primary amino groups (proteins, amino acids, DNA, phospholipids) to form uorescent products (Figure 5) (37). A uorescence assay has been successfully used to assess lipid oxidation in muscle foods and biological tissues. In addition to MA, hydroperoxides and other aldehydes also react with amino compounds generating various uorescent products with different excitation and emission maxima (37). Signicant correlations existed between this method and the TBA value as well as oxygen absorption level, and appears to be a reliable
TABLE 2. Summary of Methods for Analysis of Secondary Oxidation Products. Method TBA Compounds TBARS, mainly malonaldehyde Aldehydes, mainly alkenals Total carbonyls or specic carbonyl compound formed Volatile organic acids Comments Spectrometry technique It can be carried out on whole sample Absorption at 350-nm Standard method Spectrometry technique and HPLC for total or specic carbonyl compounds Monitoring changes in conductivity Rapid and automated Direct headspace Rapid analysis Applications All samples, especially sh oils Fats and oils Fats and oils

p-Anisidine Carbonyls

OSI method (Rancimat & Oxidative Stability Instrument) Gas Chromatography

Fats and oils

Volatile carbonyls and hydrocarbons

All samples

MEASUREMENT OF FREE RADICALS

373

indicator of oxidative deterioration in muscle foods, especially in freeze-dried products (37, 77). Table 2 exhibits the summary of methods for analysis of secondary oxidation products.

7. MEASUREMENT OF FREE RADICALS The initial steps of lipid oxidation involve chain reactions of free radicals as important short-lived intermediates. Oxidation level of fats and oils can be measured directly by detecting the formation of radicals. Methods based on the detection of radicals or on the tendency for the formation of radicals provide a good indication of initiation of lipid oxidation (78, 79). Electron spin resonance (ESR), also referred to as electron paramagnetic resonance (EPR) spectroscopy, relies on the paramagnetic properties of the unpaired electrons in radicals and has been developed for assessing the formation of free radicals originating in the early stages of oxidation and the onset of primary oxidation (6, 78). The assay measures the absorption of microwave energy when a sample is placed in a varied magnetic eld (7). Quantication of radical concentrations is complicated by comparison with stable paramagnetic compounds, such as transition metals and nitroxyl radicals (78). However, the short lifetimes and low steady-state concentration of the highly reactive lipid-derived radicals make it difcult to detect these radicals at concentrations lower than the minimum detectable concentration of 109 M (78). To overcome this problem, various approaches have been used, including pulse radiolysis and UV photolysis, continuous ow systems and spin trapping, among which spin trapping has been the most widely employed procedure (9). Spin trapping technique allows the accumulation of detectable concentrations of longer-lived radicals by addition to samples of a spin trapping agent, which reacts with free radicals to form more stable spin adducts, but often at the expense of the ability to identify the original radical (6, 9, 78). Nitroso compounds and nitrones are the most common spin traps, both leading to nitroxyl type spin adducts, such as a-phenyl-tert-butylnitrone (PBN) adducts (Figure 6) (78).
O N+ Ph CMe3 PBN + R Ph R

H N

O CMe3

O Me3C N O + R R N CMe3 MNP

Figure 6. Formation of nitroxyl radical spin adducts.

374

LIPID OXIDATION: MEASUREMENT METHODS

ESR spectroscopy is of great value for the study of the early stages of lipid oxidation and prediction of oxidative stability of fats and oils. It has high sensitivity and allows mild conditions by applying signicantly low temperatures and requires little sample preparation (6, 78, 80). Strong linear correlations were found between ESR and Rancimat and oxygen consumption analyses (6, 79). ESR has also been used for evaluation of antioxidant activity (81). Nevertheless, spin traps used in the ESR assay have been reported to exhibit widely differing trapping efciencies for different radicals and show both pro-oxidant and antioxidant effects (9, 82, 83). Moreover, spin adducts can act as antioxidants, giving erroneous results of oxidative stability of samples (9). However, even with these limitations, the ESR spectroscopy is a suitable method for measuring lipid oxidation in foods and in biological tissues.

8. OTHER METHODS 8.1. Differential Scanning Calorimetry (DSC) During lipid oxidation, fat or oil materials reveal a number of thermally induced transitions, such as the transfer of oxygen molecules to unsaturated fatty acids (exothermic process) (84). Therefore, thermal analysis can be applied in accelerated oil stability tests. The differential scanning calorimetry (DSC) technique, which is based on thermal release of oxidation reactions, has the potential as a nonchemical method for assessing oxidative stability of fats and oils, indicating the onset of advanced oxidation (termination) (6). It provides unique energy prole information, which specically measures the temperature and heat ows associated with lipid oxidation as a function of time and temperature (85). The method uses isothermal or nonisothermal conditions and a ow of oxygen as purge gas, with a calorimeter measuring the heat ow into (endothermic) or out of (exothermic) an oil sample undergoing oxidation changes (6, 84). The oxidation curves of the sample are obtained with different heating time, and a dramatic increase for the evolved heat can be observed with the appearance of a sharp exothermic curve during initiation of oxidation. The endpoint is taken at the time where a rapid exothermic reaction between oil and oxygen occurs and induction period (IP) determined automatically by intersection of extrapolated baseline and tangent line (leading edge) of the exotherm (Figure 7) (6, 84). The DSC also measures oxidation onset temperature, the temperature at maximum reaction, and the ending temperature (84). The isothermal and nonisothermal DSC show good agreement, suggesting that both isothermal and nonisothermal DSC are suitable for oxidation studies of oils (86). The DSC technique has recently been reviewed by Tan et al. (84, 87). The DSC is a sensitive, effective, and consistent method for characterization of the quality of oils at different stages of oxidation (20). It is simple and rapid, and it requires no solvent or chemical reagent. As reported by Hassel et al. (89), oils samples, which required 14 days via AOM, could be evaluated in less than 4 hours by DSC. Thus, DSC is a reliable alternative to current methods for monitoring lipid

OTHER METHODS

375

Exothermic

IP

50

100

150 Time (min)

200

250

Figure 7. Determination of induction period (IP) by DSC.

oxidation (85). The results from DSC show excellent correlations with other accelerated methods and chemical analyses (6, 73, 85). 8.2. Nuclear Magnetic Resonance (NMR) Spectroscopy High-resolution 1H NMR spectroscopy, in which hydrogen atoms (proton, 1H) with various locations in the triacylglycerol (TAG) molecules are determined, has been used to evaluate oxidative deterioration of fats and oils (7). The principle of NMR is that hydrogen atoms in a strong magnetic eld absorb energy, in the radiofrequency range, depending on their molecular environment, in which changes occur during the oxidation process (7). These changes may be monitored by NMR spectroscopy as a reection of oxidation level of food lipids. The oil sample is dissolved in CDCl3 to avoid inference from solvent, and its NMR spectrum recorded, with tetramethylsilane (TMS) as an internal standard (7). The spectrum shows several groups of signals, corresponding to the hydrogen atoms in different locations in the TAG molecules (Figure 8). The total number of each of these differently located protons can be calculated, from which ratios of aliphatic to olenic protons (Rao) and aliphatic to diallylmethylene protons (Rad) may be obtained (7). Both ratios increase steadily during lipid oxidation and may serve as an index of oxidative deterioration of oil samples. This method was reviewed by Guillen et al. (90). NMR spectroscopy has been used by many researchers, and the changes in Rao and Rad measured by NMR correlated well with Totox values (91, 92), conjugated diene values, and TBA values (93). In addition to 1H NMR, 13C NMR and 31P NMR are also powerful tools to predict oxidative stability of oils (9496). 13C NMR enables direct observation of carbon atoms. The selectivity and dispersion of 13C NMR spectra are very high (96). 13C NMR assesses lipid oxidation by monitoring the changes of carbon chains in TAG molecules, revealing the specic sites that oxidative degradation

376

LIPID OXIDATION: MEASUREMENT METHODS

g
b

H
b a b

H C O CO CH2 (CH2)n CH CH H C O CO CH2 (CH2)n CH3 H C O CO CH2 (CH2)n CH3 H

CH2 CH CH CH2 CH2 (CH2)n CH3

TMS

d e b c f

5
1

0 PPM

Figure 8. H NMR spectrum of oxidized canola oil.

occurs (94). However, because the abundance of the NMR active 13C nucleus isotope is only 1.12% of 12C, the sensitivity of 13C NMR is usually much lower than that of 1H NMR (96). NMR spectroscopy is a rapid, nondestructive, and reliable technique for assessing lipid oxidation. It simultaneously measures both the primary and the secondary oxidative changes in oils, and provides specic information on oxidative regions in the TAG molecules. Thus, NMR spectroscopy is considered a more suitable means for estimating lipid oxidation than chemical determinations. 8.3. Sensory Evaluation For the food industry, the detection of oxidative off-avors by taste or smell is the main method of deciding when a lipid-containing food is no longer t for consumption (12). Terminologies and methodologies have been developed for sensory evaluation of specic food products such as meats, peanuts, and vegetable oils (97). In the edible oil industry, the AOCS (American Oil Chemists Society) Flavor Quality Scale (revised) with separate grading and avor intensity has been employed for describing lipid oxidation (97), as summarized in Table 3. The descriptive analysis, including the detection and the description of both the qualitative and quantitative sensory aspects of a product, is performed by a trained panel, as the sensitivity to the off-avors varies among different individuals (12, 97). The sensory induction period of the product can be determined.

MEASUREMENT OF FRYING FAT DETERIORATION

377

TABLE 3. A Partial List of Terms Used to Describe Oxidized Oil. Flavor-Related Terms Buttery Nutty Beany Grassy Watermelon Painty Fishy
Adapted from (97).

Process-Oriented Terms Hydrogenated Oxidized Reverted Light-struck Rancid

Sensory evaluation of lipid oxidation has been conducted by many researchers (98100). However, as a subjective method, the reproducibility of sensory analysis is generally considered worse than that of chemical or instrumental methods. More recently, use of an electronic nose to monitor the formation of volatile compounds associated with off-avors from lipid oxidation has been proposed to supplement information from human sensory panels (101).

9. MEASUREMENT OF FRYING FAT DETERIORATION Deep-fat frying is a popular method for food preparation, in which vegetable oils not only are used as a heat-exchange medium, but also contribute to the quality of fried products (7). However, lipid oxidation easily occurs at relatively high temperatures, producing a complex series of compounds that exerts undesirable effects on food avor and quality (4). The measurement of lipid oxidation, therefore, is essential to determine its effect on food and oil quality, as well as the useful life of fats or oils subjected to frying. The oxidative changes in frying fats are characterized by a decrease in the total unsaturation of the fat with increases in the free fatty acid content, foaming, color, and viscosity as well as the content of polar compounds and polymeric material (4). Quality evaluation of frying fats, may be carried out in different ways. Physical methods estimate oxidative degradation by monitoring changes in physical properties of frying fats, such as molecular weight, specic gravity, smoke point, refractive index, chromatic parameter, viscosity, surface tension, and dielectric constant (4). Generally, rejection point of frying fat is established by sensory assessment. Chemical methods include the iodine value, saponication value, free fatty acid content, peroxide value, TBA value, or p-anisidine value, among others. PV is less useful because hydroperoxides decompose at about 150 C, and no accumulation of peroxides can be detected. The extent of oxidation can also be assessed by the analysis of oxidized fatty acids by spectroscopic means such as IR and NMR techniques (102). Moreover, GC-MS for volatile prole analysis (103) and HPLC for determination of DNPH derivatives of nonvolatile higher carbonyl compounds (62) provide qualitative

378

LIPID OXIDATION: MEASUREMENT METHODS

COOH

CH3 Double bond may present on the positions of C4, C5, C7, C8, or C9.

(CH2)nH R R = (CH2)11nCOOH

(CH2)nH R 1 < n < 11

(CH2)nH R

(CH2)nH (CH2)nH (CH2)8nCOOH n=1&2 n=3&4 (CH2)12nCOOH

Figure 9. Chemical structures of cyclic fatty acids formed during deep frying.

and quantitative evaluation of oxidation in frying fats. Cyclic fatty acids (Figure 9), which may contain hydroxy and keto groups, are formed during deep frying and can be measured by chromatography after derivatization (4, 7). Furthermore, determination of polar material in frying fats is a reliable approach for oil quality evaluation and is an ofcial method in Europe. This method involves separation of fat into a polar and nonpolar fraction via silica gel chromatography. Nonpolar fat can be weighed and the total polar material calculated or determined directly by their elution from the silica gel column (4,7). Routine analysis for frying fat deterioration has been reviewed by Gertz (104). Usually, more than two methods are required when using chemical analysis because no single group of compounds has been identied as a key indicator of oxidative degradation of frying fats.

10. METHODS FOR MEASURING ANTIOXIDANT ACTIVITY A variety of natural and synthetic antioxidants are used in fat-containing foods in order to inhibit lipid oxidation with a wide range of efciencies, depending on their properties, concentrations, and processing conditions. The need to measure antioxidant activity is well documented. Although numerous methods have been proposed for measurement of antioxidant activity, the essential features of any test are a suitable substrate, an oxidation initiator, and an appropriate measure of endpoint (9). Therefore, certain aspects should be taken into consideration when selecting a test for measuring antioxidant activity. These include the model food system used for

METHODS FOR MEASURING ANTIOXIDANT ACTIVITY

379

the test, and the means by which oxidation is accelerated and monitored (12). Normally, most assessments of antioxidant activity are performed in oil, or other model systems, giving sensible prediction for the activity in oil or water-in-oil emulsions, whereas the results may be misleading for oil-in-water emulsions (12). Furthermore, stripping of oils may be necessary in such evaluations because the endogenous antioxidants in nonstripped oils are found to enhance the oxidative stability of oils, thus giving rise to erroneous results in the efciency of antioxidants under investigation (105107). In addition to oils and fats, lipid substrates used for testing antioxidant activity could be fatty acids, fatty acid ethyl esters or triacylglycerols (9), and b-carotene (108110). In some cases, such as radical scavenging methods, no substrate is used. Most test procedures involve initiators to accelerate oxidation. The combination of increased temperature and oxygen supply, addition of metal catalysts, and exposure of the reactants to light can reduce the oxidative stability by a large amount (9, 12). Nevertheless, the elevated temperature may bring about changes in the oxidation mechanism, thus causing difculties in the prediction of
TABLE 4. Methods of Expressing Results of Antioxidant Activity Tests. Method Induction period Time to reach a set level of oxidation (preinduction period) Rate of oxidation (pre-induction period) Concentration to produce equivalent effect to reference antioxidant (pre-induction period) Concentration of ROOH functional group after set time period Concentration of oxidation product after set time period Scale reading after set time period Free stable radical quenching (DPPH) Dimensions h, d h, d mol kg1 hr1, gL1 d1 mol kg1, gL1 mequiv. kg1 mg kg1 (ppm w/w) Absorbance, conductivity, etc. Percentage inhibition EC50, concentration to decrease concentration of test free radical by 50% TEC50, time to decrease concentration of test free radical by 50% mmol peroxy radical deactivated L1 TEAC (mM Trolox equivalent to 1-mM test substance) ORAC, oxygen radical absorbance capacity; mmol of Trolox equivalents Absorbance of Fe2 complex at 593 nm produced by antioxidant reduction of corresponding tripyridyltriazine Fe3 complex Percentage of inhibition of ferrozine-Fe2 complex formation

Total radical-trapping antioxidant parameter (TRAP) ABTS assay, phycoerythrin assay Phycoerythrin assay FRAP assay

Metal chelating assay

NOTE: Also see Tables 1 and 2 for other tests applicable to antioxidant activity determination. Adapted from (9).

380

LIPID OXIDATION: MEASUREMENT METHODS

antioxidant activity at low temperatures as compared with those at high temperatures (9, 12). After the substrate is oxidized under standard conditions, the oxidation is monitored by chemical, instrumental, or sensory methods. An appropriate measure of endpoint is essential for assessing antioxidant activity. Analytical strategies for endpoint determination include measurement at a xed time point, measurement of reaction rate, lag phase measurement, and integrated rate measurement (9). The resulting antioxidant activity is expressed using a wide range of parameters (Table 4). Approaches proposed for testing antioxidant activity include measuring of the current state of oil samples, as discussed above, and radical scavenging assays, which are gaining popularity in the evaluation of antioxidant activity. Radical scavenging methods measure the relative abilities of antioxidants to scavenge synthetic radicals or natural in comparison with the antioxidant potency of a standard antioxidant compound (111). Trolox (6-hydroxy-2,5,7,8-tetramethylchroma-2-carboxylic acid), ascorbic acid, and quercetin are among the standard antioxidants frequently used. The most commonly used synthetic radicals are DPPH (2,2-diphenyl-1picrylhydrazyl) and ABTS (3-ethylbenzthiazoline-sulfonic acid) radicals. DPPH test (112116) and ABTS assay (117122) are simple, rapid, and involve no substrate. However, it has been suggested that these articial substrate-free methods do not always adequately mimic the processes in food systems, which sometimes makes them less valuable for predicting the effectiveness of the antioxidant in foods (9). Other measurements of antioxidant activity include FRAP (ferric reducingantioxidant power) (123126), TRAP (total radical-trapping antioxidant parameter) (123, 127), phycoerythrin assay (128, 129), and test of metal chelating capacity (130, 131), among others. Reviews on methods for testing antioxidant activity have been published (9, 12).

11. CONCLUSIONS AND RECOMMENDATIONS Lipid oxidation may be assessed in many ways, among which changes in the initial reactants and formation of oxidation products are most commonly assessed. Meanwhile, sensory analysis assesses both the subjective and, in some cases, objective measurements of oxidative changes in foods. Each method shows both advantages and disadvantages, thus it is important to select the most adequate method, depending on the system under investigation and the state of oxidation itself. The use of two or more methods assessing both primary and secondary oxidation products is highly recommended.

REFERENCES
1. F. Shahidi, Nahrung., 44, 158163 (2000). 2. J. R. Vercellotti, A. J. St. Angelo, and A. M. Spanier, in A. J. St. Angelo, ed., Lipid Oxidation in Food, ACS Symposium Series 500, American Chemical Society, Washington, D.C., 1992, pp. 111.

REFERENCES

381

3. M. Gordon, in J. Pokorny, N. Yanishlieva, and M. Gordon, eds., Antioxidants in Food: Practical Applications, Woodhead Publishing, Ltd., Cambridge, England, 2001, pp. 721. 4. E. G. Perkins, in A. J. St. Angelo, ed., Lipid Oxidation in Food, American Chemical Society, Washington, D.C., 1992, pp. 310321. 5. A. Kamal-Eldin, M. Makinen, and A. M. Lampi, in A. Kamal-Eldin, ed., Lipid Oxidation Pathways, AOCS Press, Champaign, Illinois, 2003, pp. 136. 6. J. Velasco, M. L. Anderson, and L. H. Skibsted, Food Chem., 85, 623632 (2004). 7. F. Shahidi and U. N. Wanasundara, in C. C. Akoh and D. B. Min, eds., Food Lipids: Chemistry, Nutrition and Biotechnology, Marcel Dekker, Inc., New York, 2002, pp. 465487. 8. M. C. Dobarganes and J. Velasco, Eur. J. Lipid Sci. Technol., 104, 420428 (2002). 9. M. Antolovich, P. D. Prenzler, E. Patsalides, S. Mcdonald, and K. Robards, Analyst, 127, 183198 (2002). 10. I. Niklova, S. Schmidt, K. Habalova, and S. Sekretar, Eur. J. Lipid Sci. Technol., 103, 299306 (2001). 11. J. Velasco and C. Dobarganes, Eur. J. Lipid Sci. Technol., 104, 661676 (2002). 12. M. Gordon, in J. Pokorny, N. Yanishlieva, and M. Gordon, eds., Antioxidants in Food: Practical Applications, Woodhead Publishing, Ltd., Cambridge, England, 2001, pp. 7184. 13. C. Genot, G. Kansci, and M. Laroche, Sciences des Aliments., 14, 673682 (1994). 14. H. J. Chung, A. S. Colakoglu, and D. B. Min, J. Food Sci., 69, 8388 (2004). 15. S. L. Melton, Food Technol., 37, 105111 (1983). 16. G. O. Fruhwirth, T. Wenzl, R. El-Toukhy, F. S. Wagner, and A. Hermetter, Eur. J. Lipid Sci. Technol., 105, 266274 (2003). 17. B. J. F. Hudson, in J. C. Allen and J. Hamilton, eds., Rancidity of Foods, Applied Science Publishers, London, England, 1983, pp. 4758. 18. A. Riuz, M. J. Ayora-Canada, and B. Lendl, Analyst, 126, 242246 (2001). 19. G. Yildiz, R. L. Wehling, and S. L. Cuppett, J. Amer. Oil Chem. Soc., 80, 103107 (2003). 20. A. Kiritsakis, A. Kanavouras, and K. Kiritsakis, Eur. J. Lipid Sci. Technol., 104, 628638 (2002). 21. Determination of Peroxide Value, in IUPAC: Standard Methods for the Analysis of Oils, Fats and Derivatives, IUPAC Standard Method 2.501, International Union of Pure and Applied Chemistry, Pergamon, Oxford, England, 1992. 22. D. L. Berner, INFORM, 1, 884886 (1996). 23. S. Eymard and C. Genot, Eur. J. Lipid. Sci. Technol., 105, 497501 (2003). 24. T. Asakawa and S. Matsushita, J. Amer. Oil. Chem. Soc., 55, 691620 (1978). 25. S. Hara and Y. Totani, J. Amer. Oil Chem. Soc., 65, 19481950 (1988). 26. M. Hicks and J. M. Gebicki, Anal. Biochem., 99, 249253 (1979). 27. J. M. Gebicki and J. Guille, Anal. Bichem., 176, 360364 (1989). 28. Z. Y. Jiang, A. C. S. Woollard, and S. P. Wolff, Lipids, 26, 853856 (1991). 29. S. P. Wolff, Methods Enzymol., 233, 182189 (1994). 30. J. M. DeLong, R. K. Prange, D. M. Hodges, C. F. Forney, M. C. Bishop, and M. Quillian, J. Agric. Food Chem., 50, 248254 (2002).

382

LIPID OXIDATION: MEASUREMENT METHODS

31. J. Sedman, F. R. van de Voort, and A. A. Ismail, in R. E. McDonald and M. M. Mossoba, eds., New Techniques and Applications in Lipid Analysis, AOCS Press, Champaign, Illinois, 1997, pp. 283324. 32. J. Dong, K. Ma, F. R. van de Voort, and A. A. Ismail, J. AOAC Int., 80, 345352 (1997). 33. T. A. Russin, F. R. van de Voort, and J. Sedman, J. Amer. Oil Chem. Soc., 80, 635641 (2003). 34. B. Innawong, P. Mallikarjunan, J. Irudayaraj, and J. E. Marcy, Lebensm. Wiss. U. Technol., 37, 2328 (2004). 35. T. Miyazawa, K. Fujimoto, and T. Kandeda, Agric. Biol. Chem., 51, 25692573 (1987). 36. A. M. Jimenez and M. J. Navas, Grasas y Aceites, 53, 6475 (2002). 37. J. P. Wold and M. Mielnik, J. Food Sci., 65, 8795 (2000). 38. C. Stauffer, in Fats & Oils, Eagan Press, St. Paul, Minnesota, 1996, pp. 1527. 39. F. Shahidi, U. Wanasundara, and N. Brunet, Food Res. Int., 27, 555562 (1994). 40. U. N. Wanasundara, F. Shahidi, and C. R. Jablonski, Food Chem., 52, 249253 (1995). 41. E. Kishida, S. Tokumaru, Y. Ishitani, M. Yamamoto, M. Oribe, H. Iguchi, and S. Kojo, J. Agric. Food Chem., 41, 15981600 (1993). 42. S. Cesa, J. Agric. Food Chem., 52, 21192122 (2004). 43. D. Jardine, M. Antolovich, P. D. Prenzler, and K. Robards, J. Agric. Food Chem., 50, 17201724 (2002). 44. A. de las Heras, A. Schoch, M. Gibis, and A. Fischer, Eur. Food Res. Technol., 217, 180 184 (2003). 45. D. V. Hoyland and A. J. Taylor, Food Chem., 40, 271291 (1991). 46. B. G. Tarladgis, B. M. Watts, M. T. Younathan, and L. Dugan, J. Amer. Oil Chem. Soc., 37, 4448 (1960). 47. F. Shahidi and C. Hong, J. Food Biochem., 15, 97105 (1991). 48. J. Pikul, D. E. Leszczynski, and F. A. Kummerow, J. Agric. Food Chem., 37, 13091313 (1989). 49. C. Wang, L. Zhu, and M. S. Brewer, in 1996 IFT annual meeting: Book of Abstracts, pp. 161. 50. R. Guillen-Sans and M. Guzman-Chozas, Crit. Rev. Food Sci. Nutr., 38, 315330 (1998). 51. Q. Sun, C. Faustman, A. Senecal, A. L. Wilkinson, and H. Furr, Meat Sci., 57, 5560 (2001). 52. F. Shahidi, L. J. Rubin, L. L. Diosady, and D. F. Wood, J. Food Sci., 55, 274275 (1985). 53. R. Marcuse and J. Pokorny, Fett. Wiss. Technol., 96, 185187 (1994). 54. H. de A. Gomes, E. N. da Silva, M. R. Lopes do Nascimento, and H. T. Fukuma, Food Chem., 80, 433437 (2003). 55. F. Doleschall, Z. Kemeny, K. Recseg, and K. Kovari, Eur. J. Lipid Sci. Technol., 104, 14 18 (2002). 56. G. H. van der Merwe, L. M. du Plessis, and J. RN. Taylor, J. Sci. Food Agric., 84, 5258 (2003). 57. G. R. List, C. D. Evans, W. F. Kwolek, K. Warner, and B. K. Boundy, J. Amer. Oil Chem. Soc., 51, 1721 (1974). 58. M. D. Guillen and N. Cabo, Food Chem., 77, 503510 (2002).

REFERENCES

383

59. P. J. White, in K. Warner and N. A. M. Eskin, eds., Methods to Assess Quality and Stability of Oils and Fat-containing Foods, AOCS Press, Chamaign, Illinois, 1995, pp. 159178. 60. U. N. Wanasundara and F. Shahidi, J. Food Lipids, 2, 7386 (1995). 61. T. Chiba, M. Takazawa, and K. Fujinoto, J. Amer. Oil Chem. Soc., 66, 15881592 (1989). 62. E. Schulte, Anal. Bioanal. Chem., 372, 644648 (2002). 63. F. Shahidi and R. B. Pegg, J. Food Lipids, 1, 177186 (1994). 64. C. F. Goodridge, R. M. Beaudry, J. J. Pestka, and D. M. Smith, J. Agric. Food Chem., 51, 41854190 (2003). 65. H. Z. Zhang and T. C. Lee, IFT Annual Meeting, 1995, pp. 161. 66. C. F. Goodridge, R. M. Beaudry, J. J. Pestka, and D. M. Smith, J. Agric. Food Chem., 51, 75337539 (2003). 67. U. N. Wanasundara and F. Shahidi, Paper presented at the Canadian Section of American Oil Chemists Societys Annual Meeting, Guelph, Ontario, Canada, November 1516, 1995. 68. F. Shahidi and S. A. Spurvey, J. Food Lipids, 3, 1325 (1996). 69. K. Schwarz, G. Bertelsen, L. R. Nissen, P. T. Gardner, M. I. Heinonen, A. Hopia, T. Huynh-Ba, P. Lambelet, D. McPhail, L. H. Skibsted, and L. Tijburg, Eur. Food Res. Technol., 212, 319328 (2001). 70. K. Miura, H. Kikuzaki, and N. Nakatani, J. Agric. Food Chem., 50, 18451851 (2002). 71. S. de la Presa-Owens, M. C. Lopez-Sabater, and M. Rivero-Urgell, J. Agric. Food Chem., 43, 28792882 (1995). 72. M. P. Canizares-Macias, J. A. Garcia-Mesa, and M. D. Luque de Castro, Anal. Bioanal. Chem., 378, 479483 (2004). 73. C. P. Tan, Y. B. Che-Man, J. Selamat, and M. S. A. Yusoff, Food Chem., 76, 385389 (2002). 74. R. Aparicio, L. Roda, M. A. Albi, and F. Gutierrez, J. Agric. Food Chem., 47, 41504155 (1999). 75. A. Faroop, M. I. Bhanger, and T. G. Kazi, J. Amer. Oil Chem. Soc., 80, 151155 (2003). 76. L. C. Boyd, V. C. Nwosu, C. L. Young, and L. MacMillian, J. Food Lipids, 5, 269282 (1998). 77. A. L. Wilkinson, Q. Sun, A. Senecal, and C. Faustman, J. Food Sci., 66, 2024 (2001). 78. M. L. Andersen and L. H. Skibsted, Eur. J. Lipid Sci. Technol., 104, 6568 (2002). 79. C. U. Carlsen, M. L. Andersen, and L. H. Skibsted, Eur. Food Res. Technol., 213, 170 173 (2001). 80. M. K. Thomsen, D. Kristensen, and L. H. Skibsted, J. Amer. Oil Chem. Soc., 77, 725730 (2000). 81. L. R. Nissen, Tuong-Huynh-Ba, M. A. Petersen, G. Bertelsen, and L. H. Skibsted, Food Chem., 79, 387394 (2002). 82. B. Halliwell and J. M. C. Gutteridge, in Free Radicals in Biology and Medicine, Oxford University Press, New York, 1999. 83. D. M. Lee, Methods Enzymol., 234, 513523 (1994). 84. C. P. Tan and Y. B. Che Man, Trends Food Sci. Technol., 13, 312318 (2002). 85. C. P. Tan and Y. B. Che Man, Food Chem., 67, 177184 (1999).

384

LIPID OXIDATION: MEASUREMENT METHODS

86. G. Litwinienko, A. Daniluk, and T. Kasprzycka-Guttman, J. Amer. Oil Chem. Soc., 76, 655657 (1999). 87. C. P. Tan and Y. B. Che Man, Lipid Technol., 14, 112115 (2002). 88. B. Kowalski, K. Ratusz, D. Kowalska, and W. Bekas, Eur. J. Lipid Sci. Technol., 106, 165169 (2004). 89. R. L. Hassel, J. Amer. Oil Chem. Soc., 53, 179181 (1976). 90. M. D. Guillen and A. Ruiz, Trends Food Sci. Technol., 12, 328338 (2001). 91. F. Shahidi, U. Wanasundara, and N. Brunet, Food Res. Int., 27, 555562 (1994). 92. U. Wanasundara and F. Shahidi, J. Food Lipids, 1, 1524 (1993). 93. S. P. J. Namal-Senanayake and F. Shahidi, J. Food Lipids, 6, 195203 (1999). 94. I. Medina, R. Sacchi, I. Giudicianni, and S. Aubourg, J. Amer. Oil Chem. Soc., 75, 147 154 (1998). 95. F. J. Hidalgo, G. Gomez, J. L. Navarro, and R. Zamora, J. Agric. Food Chem., 50, 5825 5831 (2002). 96. B. W. L. Diehl, in R. J. Hamilton, ed., Lipid Analysis in Oils and Fats, Blackie Academic & Professional, London, 1998, pp. 87135. 97. G. V. Cinille and C. A. Dus, in A. J. St. Angelo ed., Lipid Oxidation in Food, American Chemical Society, Washington, D.C., 1992, pp. 279289. 98. C. J. Broadbent and O. A. Pike, J. Amer. Oil Chem. Soc., 80, 5963 (2003). 99. E. A. Coppin and O. A. Pike, J. Amer. Oil Chem. Soc., 78, 1318 (2001). 100. M. Timm-Heinrich, X. Xu, N. S. Nielsen, and C. Jacobsen, Eur. J. Lipid Sci. Technol., 105, 436448 (2003). 101. N. Shen, S. Moizuddin, L. Wilson, S. Duvick, P. White, and L. Pollak, J. Amer. Oil Chem. Soc., 78, 937940 (2001). 102. S. Khatoon and A. G. G. Krishna, J. Food Lipids, 5, 247267 (1998). 103. V. Rosales and J. de D, Dissertation Abstracts International, B., 57, 7290 (1997). 104. C. Gertz, Lipid Technol., 13, 4447 (2001). 105. T. Keceli and M. H. Gordon, J. Sci. Food Agric., 81, 13911396 (2001). 106. M. A. Khan and F. Shahidi, J. Amer. Oil Chem. Soc., 77, 963968 (2000). 107. V. Povilaityte, M. E. Cuvelier, and C. Berset, J. Food Lipids, 8, 4564 (2001). 108. F. Shahidi, C. Desilva, and R. Amarowicz, J. Amer. Oil Chem. Soc., 80, 443450 (2003). 109. F. Shahidi and R. Amarowicz, J. Amer. Oil Chem. Soc., 73, 11971199 (1996). 110. R. Amarowicz, M. Naczk, and F. Shahidi, J. Amer. Oil Chem. Soc., 77, 957961 (2000). 111. O. A. Zaporozhets, O. A. Krushynska, N. A. Lipkovska, and V. N. Barvinchenko, J. Agric. Food Chem., 52, 2125 (2004). 112. D. Bandoniene, M. Murkovic, W. Pfannhauser, P. R. Venskutonis, and D. Gruzdiene, Eur. Food Res. Technol., 214, 143147 (2002). 113. N. Nenadis and M. Tsimidou, J. Amer. Oil Chem. Soc., 79, 11911195 (2002). 114. H. Matsuura, H. Chiji, C. Asakawa, M. Amano, T. Yoshihara, and J. Mizutani, Biosci. Biotechnol. Biochem., 67, 23112316 (2003). 115. P. J. Park, F. Shahidi, and Y. J. Jeon, J. Food Lipids, 11, 1528 (2004). 116. R. Amarowicz, B. Raab, and F. Shahidi, J. Food Lipids, 10, 5162 (2003).

REFERENCES

385

117. A. Podsedek, D. Sosnowska, and B. Anders, Eur. Food Res. Technol., 217, 296300 (2003). 118. A. M. Alonso, D. A. Guillen, and C. G. Barroso, Eur. Food Res. Technol., 216, 445448 (2003). 119. N. J. Miller and C. Rice-Evan, Redox Rep., 2, 161171 (1996). 120. C. E. Rice-Evans and N. J. Miller, Methods Enzymol., 234, 279 (1994). 121. L. Bramati, F. Aquilano, and P. Pietta, J. Agric. Food Chem., 51, 74727474 (2003). 122. N. J. Miller and C. A. Rice-Evans, Free Rad. Res., 26, 195199 (1997). 123. N. Pellegrini, M. Serani, B. Colobi, D. del Rio, S. Salvatore, M. Bianchi, and F. Brighenti, J. Nutr., 133, 28122819 (2003). 124. I. E. F. Benzie and J. J. Strain, Anal. Biochem., 239, 70 (1996). 125. I. E. F. Benzie and J. J. Strain, Methods Enzymol., 299, 15 (1999). 126. J. Chen, H. Lindmark-Mansson, L. Gorton, and B. Akesson, Int. Dairy J., 13, 927935 (2003). 127. A. Ghiselli, M. Serani, G. Maiani, E. Azzini, and A. Fero-Luzzi, Free Rad. Biol. Med., 18, 2936 (1995). 128. O. Boxin, M. Hampsch-Woodill, and R. L. Prior, J. Agric. Food Chem., 49, 46194626 (2001). 129. S. McDonald, P. D. Prenzler, M. Antolovich, and K. Robards, Food Chem., 73, 7384 (2001). 130. S. N. El and S. Karakaya, Int. J. Food Sci. Nutr., 55, 6774 (2004). 131. M. Oktay, I. Gulcin, and O. I. Kufrevioglu, Lebensm. Wiss. U. Technol., 36, 263271 (2003).