Food Chemistry 145 (2014) 918–926

Contents lists available at ScienceDirect

Food Chemistry

journal homepage: www.elsevier .com/locate / foodchem

Review

Sample preparation: A critical step in the analysis of cholesteroloxidation products

0308-8146/$ - see front matter � 2013 Elsevier Ltd. All rights reserved.http://dx.doi.org/10.1016/j.foodchem.2013.08.123

⇑ Corresponding author. Tel.: +357 22 892774; fax: +357 22 892801.E-mail address: ckapni1@ucy.ac.cy (C.P. Kapnissi-Christodoulou).

1 These authors contributed equally to this work.

Christiana A. Georgiou 1, Michalis S. Constantinou 1, Constantina P. Kapnissi-Christodoulou ⇑Department of Chemistry, University of Cyprus, 1678 Nicosia, Cyprus

a r t i c l e i n f o

Article history:Received 4 March 2013Received in revised form 28 June 2013Accepted 28 August 2013Available online 7 September 2013

Keywords:Cholesterol oxidation productsLipid extractionSample preparationSaponificationSolid phase extraction

a b s t r a c t

In recent years, cholesterol oxidation products (COPs) have drawn scientific interest, particularly due totheir implications on human health. A big number of these compounds have been demonstrated to becytotoxic, mutagenic, and carcinogenic. The main source of COPs is through diet, and particularly fromthe consumption of cholesterol-rich foods. This raises questions about the safety of consumers, and itsuggests the necessity for the development of a sensitive and a reliable analytical method in order toidentify and quantify these components in food samples. Sample preparation is a necessary step in theanalysis of COPs in order to eliminate interferences and increase sensitivity. Numerous publications have,over the years, reported the use of different methods for the extraction and purification of COPs. How-ever, no method has, so far, been established as a routine method for the analysis of COPs in foods. There-fore, it was considered important to overview different sample preparation procedures and evaluate thedifferent preparative parameters, such as time of saponification, the type of organic solvents for fatextraction, the stationary phase in solid phase extraction, etc., according to recovery, precision andsimplicity.

� 2013 Elsevier Ltd. All rights reserved.

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9182. Lipid extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9203. Saponification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9214. Solid phase extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9235. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 925

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 925

1. Introduction

Cholesterol is vulnerable to oxidation, leading to the formationof a variety of cholesterol oxidation products, the so-called COPs. Inrecent years, COPs have gained considerable attention due to theirimplications in human health (Clariana, Díaz, Sárraga, & García-Regueiro, 2011a). These compounds have been well documentedfor being potentially cytotoxic, mutagenic, carcinogenic, and theyhave been associated with the promotion of atherosclerosis(Brown & Jessup, 1999; Leonarduzzi, Sottero, & Poli, 2002; Lordan,Mackrill, & O’Brien, 2009; Olkkonen, Béaslas, & Nissilä, 2012;

Otaegui-Arrazola, Menendez-Carreño, Ansorena, & Astiasarán,2010; Poli, Sottero, Gargiulo, & Leonarduzzi, 2009). It is thoughtthat COPs modulate the structure and function of the cellularmembrane and inhibit the activity of enzymes involved in choles-terol biosynthesis (Bielska, Schlesinger, Covey, & Ory1, 2012;Brown & Jessup, 2009; Gill, Chow, & Brown, 2008; Jusakul, Yongva-nit, Loilome, Namwat, & Kuver, 2011; Olsen, Schlesinger, & Baker,2009; Olsen, Schlesinger, Ory, & Baker, 2012).

The main source of COPs is through diet, and particularly fromthe consumption of cholesterol-rich foods, such as meat, eggs anddairy products (Sampaio, Bastos, Soares, Queiroz, & Torres, 2006;Vicente & Torres, 2007). It is generally accepted that fresh foodscontain very low levels of COPs (Cardenia, Rodriguez-Estrada, Bald-acci, Savioli, & Lercker, 2012). However, inadequate storage, cook-ing and processing conditions tend to increase the degree of

Fig. 1. Structural formulas of the most important COPs and cholesterol.

C.A. Georgiou et al. / Food Chemistry 145 (2014) 918–926 919

cholesterol oxidation (Echarte, Ansorena, & Astiasarán, 2003; Hur,Park, & Joo, 2007; Olkkonen et al., 2012; Pignoli et al., 2009; Sa-vage, Dutta, & Rodriguez-Estrada, 2002). This, therefore, raisesquestions about the safety of the consumers, and it suggests thatit is imperative to develop a sensitive and a reliable analyticalmethod in order to monitor these components in food samples(Yen, Inbaraj, Chien, & Chen 2010). The most common COPs foundin several foodstuffs are demonstrated in Fig. 1.

Gas chromatography (GC) and high performance liquid chroma-tography (HPLC) are the most widely used techniques for the anal-ysis of COPs. The former can provide better resolution, but itrequires a time-consuming derivatization of the analytes in orderto enhance their volatility and thermal stability (Chen, Chien,Inbaraj, & Chen, 2012). HPLC, which is considered as the alternativemethod to GC, enables the analysis of these compounds in rela-tively low temperatures without the need for derivatization (Raithet al., 2005; Vicente, Sampaio, Ferrari, & Torres, 2012). Hence, it is afast and a simple analytical methodology. The most commondetection systems used for the qualitative and quantitative deter-mination of COPs are spectrophotometric (UV), refractive index(RI), flame ionization (FI) and mass spectrometric (MS). Most ofCOPs absorb at low wavelengths (200–215 nm region), where mostorganic compounds exhibit some UV absorbance. This, in turn,makes the UV detector less selective. In addition, a UV detector isnot useful for the detection of some biological important oxyster-ols, such as 5,6-epoxy-cholesterol or cholestane-3b,5a,6b-triol,which do not possess adequate UV absorption characteristics(Saldanha, Sawaya, Eberlin, & Bragagnolo, 2006; Vicente et al.2012). The use of MS and tandem MS/MS opens access to moreselective detection, reducing matrix background and improving

profile with a low S/N ratio (Clariana et al., 2011; Saldanha et al.,2006).

Prior to chromatographic determination and quantification ofCOPs, sample preparation is necessary to be employed in order toeliminate interferences and increase sensitivity. Although numer-ous research articles have, over the years, been published concern-ing extraction and purification of COPs from foodstuffs, no certifiedmethod is yet available. Thus, the development of a credible assayis an urgent problem that needs to be overcome (Chen et al., 2012;Janoszka, 2010; Sieber, 2005).

This particular step of the analytical protocol is considered to bethe most challenging for several reasons. The fact that COPs arepart of the complex mixture of lipids, where they are mostly pres-ent in trace levels (ppm or ppb) indicating that the extensive work-up and cleaning procedures before final quantification are re-quired. However, one major concern that needs special attentioninvolves the possibility of generation of the so-called artifacts (Car-denia et al., 2012; Lozada-Castro, Gil-Diaz, Santos-Delgado, Rubio-Barroso, & Polo-Diez, 2011; Manini, Andreoli, Careri, Elviri, & Mus-ci, 1998). Artifacts are COPs that are formed under analytical con-ditions, either due to cholesterol oxidation or due to the conversionof pre-existing COPs to others (Busch & King, 2009). The bestknown example is the degradation of 7-keto to cholesta-3,5-dien-7-one in alkaline medium (Lozada-Castro et al., 2011). Someauthors in order to limit artifact generation, recommended theuse of antioxidants, such as butylated hydroxytoluene (BHT), dur-ing sample preparation (Janoszka, 2010; Du, Nam, & Ahn, 2001;Nam, Du, Jo, & Ahn, 2001).

The general route in COPs analysis involves three major steps:extraction of lipids from the food matrix, hydrolysis of esterified

920 C.A. Georgiou et al. / Food Chemistry 145 (2014) 918–926

sterols by saponification and subsequent enrichment of COPs withsolid phase extraction (SPE) (Sieber, 2005; Ubhayasekera, Verleyen,& Dutta, 2004). However, this is not a standard procedure, as manystudies use some modifications (Table 1). The objective of thisstudy is the extensive presentation of the main existing approachesof each analytical step mentioned above.

2. Lipid extraction

Lipids exist in biological matrices mainly in two major forms;they are present as droplets in storage tissues or they can be con-stituents of cell membranes (Christie, 1993; Farese & Walther,2009; Guardiola, Dutta, Codony, & Savage, 2002; Reue, 2011). Ineach case, they are not only closely associated with each other,but they interact with non-lipid material, such as proteins, viahydrophobic and Van der waals forces, hydrogen bonding and elec-trostatic interactions, as well (Christie, 1993; Jensen, & Mouritsen,2004). For this reason, the isolation of liposoluble compounds fromfoodstuff, including COPs, is not easy, and thus, the use of a methodcapable to provide total extraction of lipids is necessary for a reli-able analysis. In principle, the quantitative extraction of lipids fromtissues can be achieved with the use of an ideal solvent, whichcould simultaneously dissolve the lipids and disrupt the polarand hydrophobic interactions between them and other tissue ma-trix (Iverson, Lang, & Cooper, 2001; Guardiola et al., 2002). There-fore, the use of a single apolar organic solvent, such as hexane, isunsuitable. To meet this requirement, a fairly polar solvent or mix-ture must be employed (Christie, 1993; Iverson et al. 2001).

Folch, Lees, and Stanley (1957) were among the first to recog-nize this aspect, and they developed a method using a mixture ofchloroform/methanol (2:1, v/v) for the lipid extraction. Regardingthe proper amount of sample and solvents, they suggested that tis-sue should be homogenized with chloroform/methanol (2:1, v/v)to a final volume 20 times the volume of the tissue sample. Dueto the toxicity of chloroform, alternative solvent systems have beendeveloped. A method described by Hara and Radin involved the useof n-hexane/2-propanol (3:2, v/v) mixture as the lipid extractant,while Maxwell’s method reported the use of dichloromethaneand methanol (Hara & Radin, 1978; Maxwell, Mondimore, & To-bias, 1986). Apart from a re-development of Hara and Radin’smethod, these two procedures have not found wide applicationin COPs analysis (Ferioli, Caboni, & Dutta, 2008; Ferioli, Dutta, &Caboni, 2010; Ubhayasekera, Tres, Cobony, & Dutta, 2010). In brief,Folch’s method is by far the most widely used method, as itcontinues to be considered the most efficient and reliable tech-nique for the exhaustive isolation of lipids from various tissues

Table 1Different pathways of sample preparation in COPs analysis.

Matrix Lipid extraction* Purification

Porcine patties Chloroform/methanol (1:2,v/v) Si-SPE & NHDry-cured shoulder Methanol/chloroform/water

(2:1:1, v/v)Si-SPE & NH

Pig feet meat and skin Chloroform/hexane (2:1,v/v) Si-SPETea-leaf eggs Chloroform/methanol (2:1,v/v) Si-SPECooked ham, Serrano ham, minced

beef, and soft cheeseChloroform Si-SPE

Minced beef n-Hexane/i-propanol (3:2,v/v) Cold sapon1 M KOH in

Pickled mackerel Dichloromethane/methanol(2:1,v/v)

Cold saponwith 1 M K

Pates and pork sausage Chloroform/methanol (2:1,v/v) Cold sapon2 N NaOH i

Deep-fried pork rinds, dried beef andsun-dried shrimp

Chloroform/methanol (1:1,v/v) Cold sapon1 M KOH in

* Lipid extraction referred to the first solvent system used for sample homogenization.

(Grau, Cobony, Grimpa, Baucells, & Guardiola 2001; Janoszka,2010; Mazalli & Bragagnolo, 2007; Mazalli & Bragagnolo, 2009;Obara, Obiedzinski, & Kołczak, 2006; Sampaio et al. 2006). Inevita-bly, the use of a quite polar solvent system will lead to co-extrac-tion of non-lipid material; hence, Folch et al. suggested anappropriate washing step in order to remove these contaminantsbefore proceeding to the next step of the analytical protocol,which, in our case, it is the purification of COPs. This step is essen-tial in order to partition the system into two phases; the lower or-ganic layer containing all of the tissue lipids and the upper oneconsisting of non-lipid co-extracted contaminants.

However, one important parameter, which can influence thequantitative recovery of the more polar lipids, such as COPs, isthe amount of water or salt solution that is added after samplehomogenization with chloroform/methanol (2:1, v/v). Accordingto the original paper, the proposed volume ratio of chloroform/methanol/water must be exactly 8:4:3. These proportions shouldbe constant; otherwise, selective losses of these lipids, into upperphase, may occur (Christie, 1993; Guardiola et al. 2002). Neverthe-less, it seems that this critical aspect is often overlooked in somestudies (Chen, Lu, Chien, & Chen, 2010; Lee, Chien, & Chen 2006;Lee, Chien, & Chen 2008).

Several slightly modified versions of Folch’s method also exist,with the version proposed by Boselli, Velazco, Caboni, and Lerc-ker, (2001) being the most known to COP research society. Partic-ularly, the samples were homogenized with chloroform/methanol(1:1, v/v), and they were kept at 60 �C in order to maximizeextraction recovery yield. The mixture was re-homogenized withan equal amount of chloroform, and then, it was passed throughfilter paper to eliminate the solid residue, which mainly consistedof proteins. Subsequently, the filtrate, which contained the lipidsaccompanied by non-lipid substances, was mixed with 1 M KClsolution and was left overnight at 4 �C for phase separation. Thelower phase, freed from non-lipid substances, was collected anddried with a vacuum evaporator for further purification of COPs(Boselli et al., 2005, Bonoli, Caboni, Rodriguez-Estrada, & Lercker,2007; Soto-Rodriguez et al., 2008; Boselli, Rodriguez-Estrada, Fed-rizzi, & Caboni, 2009; Verardo et al., 2010; Bosselli, Rodriguez-Estrada, Ferioli, Caboni, & Lercker, 2010; Cardenia et al., 2011;Pignoli et al. 2009). In our knowledge, no studies were performedby the authors to clarify whether the temperature could induceartifact formation.

The second most known lipid extraction procedure was devel-oped by Bligh and Dyer (Bligh & Dyer, 1959). This method is stillused as an alternative to Folch’s method, but to a lesser extent inCOPs analysis (Broncano, Petrón, Parra, & Timón, 2009; Clariana& García-Regueiro, 2011b; Clariana et al., 2011a; Petrón,

Enrichment References

2-SPE Rodríguez-Carpena et al. (2012)2-SPE Clariana et al. (2011a)

Chen et al., (2011)Chen et al. (2010)Lozada-Castro et al. (2011)

ification (RT, 18 h, withEtOH)

2-foldSi-SPE

Ferioli et al. (2008)

ification (RT, 18–20 h,OH in EtOH)

Si-SPE Ubhayasekera, Jayasinghe,Ekanayake, and Paresh (2012)

ification (RT, 1–2 h, withn MeOH)

Derewiaka and Obiedzinski (2010)

ification (RT, 20 h, withMeOH)

NH2-SPE Soto-Rodriguez et al. (2008)

C.A. Georgiou et al. / Food Chemistry 145 (2014) 918–926 921

Garcia-Regueiro, Martin, Muriel, & Antequera, 2003; Rodriguez-Carpena, Morcuende, Petrón, & Estevez, 2012). As mentionedearlier, the water content withheld in the tissue is taken into ac-count as a ternary component of the extraction system. Initially,the sample, containing endogenous water, is homogenized with amixture of chloroform/methanol (1:2, v/v), resulting in the forma-tion of a monophasic solution. After that, further homogenizationtakes place with chloroform and water, leading to a biphasic sys-tem. The addition of chloroform is necessary in order to ensurethe complete extraction of lipids. The upper phase is discarded,while the chloroform layer is concentrated for the isolation ofCOPs. The suggested proportions for a sample, containing 80%endogenous water, after water and chloroform addition, is2:2:1.8 for chloroform, methanol and water, respectively. In cases,where the moisture content is less than 80%, it is necessary eitherto add water or reduce solvent volumes in order to meet therequirements described above (Bligh and Dyer, 1959).

Although there are very few reports available regarding thecomparison between different extraction methods, they haveclearly demonstrated that the existing ones are not totally equiva-lent in their extraction efficiency, and therefore, they must be eval-uated prior to application. Chen et al. (2010) evaluated differentsolvent systems in order to enhance COPs extraction efficiency inegg samples. Various solvents, including hexane, methanol, ethylacetate, acetone and chloroform were examined either separatelyor combined. The mixture of chloroform/methanol (2:1 v/v) provedto be the most suitable because it minimized the amount of impu-rities in eggs. In contrast, Chen et al. (2012) followed the same pro-cedure for the analysis of pig feet samples, and they observed thatthis system was less appropriate as it led to coextraction of inter-fering compounds. Better results were obtained when chloroform/hexane (2:1, v/v) was adopted. These two studies strengthened theview, which suggests that the selection of a particular solvent sys-tem is affected by the nature of the food matrix.

Even though the use of a single solvent for lipid extraction israre, Lozada-Castro et al. (2011), comparing different sample prep-aration procedures, indicated that the use of pure chloroform wasthe optimum choice. Moreover, the authors validated the methodin terms of extraction time (6, 12, 18 h) and solvent volume (40,70, 100 ml). The evaluation data demonstrated that long extractiontime resulted in better recoveries, while the volumes of 70 and100 ml provided comparable results. In a recent study, Georgiou& Kapnissi-Christodoulou, 2013 compared and evaluated twoextraction solvent systems, chloroform/methanol (2:1 v/v) and n-hexane/2-propanol (3:2 v/v). The former proved to be superiorwith a very good recovery value (106 ± 8%), while the latter systemprovided a dramatic decrease of recovery (20 ± 4%). One possibleexplanation was the quite apolar nature of solvents that were usedin fat extraction (Georgiou & Kapnissi-Christodoulou, 2013). Thisobservation was in agreement with Dionisi et al., who also notedthat Folch’s method generated the least percentage of artifactscompared to Radin’s and Maxwell’s (Dionisi, Golay, Aeschlimann,& Fay, 1998).

Fig. 2. Degradation of 7-keto during hot saponification.

3. Saponification

Purification is the most critical step of the analytical procedure,since it demands selective isolation of the COPs from lipid fraction(Ruiz-Gutierrez & Perez-Camino, 2000). This ‘‘clean-up’’ step hasbeen challenging for some decades since COPs are present in minorquantities in several foodstuffs (Tai, Chen, & Chen, 1999). If we alsoconsider that more than 95% of dietary fat is in the form oftriacylglycerols and the remaining part consists of phospholipids,esterified and free cholesterol, partial glycerides, free fatty acidsand lipid-soluble vitamins, their isolation from this complex lipid

mixture is even more difficult (Ulberth and Rossler, 1998). Anefficient purification procedure should, therefore, remove COPsfrom the bulk of lipids, as these compounds, if they contaminatethe isolated COP containing fraction can interfere with COPs quan-tification (Cardenia et al. 2012; Chen et al. 2010). Saponificationand/or SPE are usually employed in order to increase their relativeconcentration prior to chromatographic analysis.

Saponification (alkaline hydrolysis) plays two important rolesin the context of COPs analysis. Firstly, it removes the predominat-ing acylglycerols of the crude lipid extract by converting them intowater-soluble soaps and free glycerol, and secondly, it hydrolyzescholesterol esters. In the saponification procedure, lipid extract isincubated in methanolic or ethanolic solution of NaOH or KOHfor a period of time, and this is followed by liquid/liquid extractionfor the concentration of the non-saponifiable fraction (Guardiolaet al. 2002). Much of the research regarding saponification has fo-cused on artifact generation. The temperature of incubation andthe alkalinity concentration were recognized as two critical param-eters, which resulted in artifact formation by degrading unstableCOPs, e.g. 7-ketocholesterol (Fig. 2) (Busch & King, 2009).

In a recent study, Busch et al. investigated the stability of 7-ketocholesterol in solution at various temperatures during sapon-ification. Particularly, 7-keto was subjected to 1 M methanolicKOH at 37 �C for 18 h or 45 �C for 3 h, and the recovery results werecompared to that obtained under control conditions at 24 �C for18 h. At temperatures greater than 24 �C, the recovery of 7-ketowas only 53% (37 �C) and 49% (45 �C) in relation to the control, sug-gesting sensibility at high temperatures. These findings also indi-cated that the temperature of saponification affected at a greaterextent the degradation in respect to the time of incubation. Thesame group reported further work on the stability of 7-keto in dif-ferent alkaline conditions (3.6 M KOH for 3 h and 1 M KOH for 18 at24 �C). When the sample was treated with a stronger alkaline solu-tion, the recovery dropped to 71%, revealing the liability of 7-keto(Busch & King, 2010). These observations are in agreement with acorresponding study performed by Park, Guardiola, Park, and Addis(1996) who suggested that 7-keto was sensitive to minor changesin temperature and it was also influenced by the strength of thealkaline reagents that were used.

However, it should be pointed out that the use of saponificationis not prohibitive. As suggested by many researchers, if saponifica-tion is conducted at room temperature under mild conditions (1 MKOH), it can be considered safe with minimal artifact generation(Tai et al. 1999). For instance, Park et al. (1996) reported that care-ful implementation of cold saponification led to 97% recovery of 7-keto, and thus, the degradation was practically negligible. Consid-ering the above, many saponification protocols have been createdat these conditions (Ferioli et al. 2008; Soto-Rodriguez et al.2008; Cardenia et al. 2012; Baggio & Bragagnolo, 2006; Baggio,Miguel, & Bragagnolo, 2005; Bonoli et al. 2007; Morales-Aizpurua& Tenuta-Filho, 2005).

In a method comparison involving three different saponificationmethods, the criteria for the selection of the optimum one were therecovery data obtained for each COP at three concentration levels(5, 10 and 20 lg) (Ubhayasekera et al. 2004). Methods A and B,were performed at room temperature for 18 h, but they differed

922 C.A. Georgiou et al. / Food Chemistry 145 (2014) 918–926

from each other, mainly on the type of solvent used in alkalinehydrolysis. In the former method, 95% ethanol was used, while inthe latter, 95% methanol in water was used. In method C, the sam-ple was incubated in ethanolic solution at 60 �C for 1 h. Method Ademonstrated the highest recoveries at all levels of spiked COPs.This indicated that this procedure was the most efficient for theCOPs isolation from tallow. Therefore, the solvent of the alkalinesolution can even affect the final concentration. As expected, hotsaponification provided the lowest recovery of 7-keto when com-pared to the other two methods. This enhanced the above notionthat the degradation of COP occurred under thermal conditions.

On the other hand, many investigators, in order to circumventthe problem of artifact generation, replaced the saponification stepwith SPE in order to isolate and enrich COPs. SPE is a faster and amilder technique in relative to saponification (Chen et al. 2010;Chen et al. 2012; Ulberth and Rossler, 1998). Purification by SPEwill be discussed later.

Direct saponification of the food matrix, without prior lipid iso-lation, is a viable option for sample preparation (Mariutti, Nogue-ira, & Bragagnolo, 2008; Mazalli, Sawaya, Eberlin, & Bragagnolo2006; Saldanha, Benassi, & Bragagnolo, 2008; Saldanha & Bragag-nolo, 2007; Saldanha et al. 2006). In a study by Saldanha et al.(2006), sample preparation was carried out using direct saponifica-tion, followed by extraction of the unsaponifiable material. Theinvestigation of different saponification factors, including percent-age of KOH in water (20–50%), volume of ethyl alcohol (2–6 mL)and time of saponification (12–24 h), demonstrated that theseparameters play a crucial role for reliable final quantification. Totaldissolution of the sample and prevention of emulsion wereachieved by using a solution of 50% KOH in water added to 6 mLof ethyl alcohol for 22 h. Furthermore, they examined the type of

Fig. 3. SPE elution systems examined by Guardiola et al. (199

solvent (n-hexane and diethyl ether) and the number of extrac-tions (three, four and five) of the nonsaponifiable matter in orderto optimize the methodology of saponification. N-hexane was cho-sen as the ideal extraction solvent over diethyl ether due to higherrecoveries. The analysis data of samples, which underwent fourand five successive extractions, indicated no significant differ-ences; hence, four extractions were selected. Similar results wereobtained by Mariutti et al. (2008), who supported that hexanewas the most appropriate extraction solvent, since fewer interfer-ence peaks were observed.

One interesting study, which was cited in numerous researcharticles and reviews, was carried out by Dionisi et al. (1998). Theycompared four of the major extraction procedures used for COPsanalysis in milk powders. Three of them involved a preliminaryfat extraction (Folch’s, Radin’s and Maxwell’s method) followedby a saponification step, while the last one involved a direct sapon-ification step. Each method performance was evaluated accordingto the following four variables: standard deviation, coefficient ofvariation, artifact generation, and recovery. The direct methoddemonstrated the best compromise among the four alternatives,because it provided good repeatability and the lowest artifact for-mation. Furthermore, this procedure was superior to others interms of time analysis and solvent quantity.

However, the application of direct saponification in more com-plex matrices may be unsuitable (Busch & King, 2009). Lozada-Cas-tro et al. (2011) observed that direct saponification, which wasapplied on cooked ham, produced emulsions that were difficultto handle. In addition, Georgiou & Kapnissi-Christodoulou, 2013 re-ported that the use of this method in meat samples provided chro-matograms with interfering peaks, making the identification of22R-hydroxycholesterol impossible.

5) for the enrichment of COPs. (⁄ COP extraction solvent).

Table 2The most frequently used SPE protocols with schematic representation.

COPs

e

f

g

e, f apolar lipids & cholesterolg

COPs

a

b

c

d

a, b, c

d

apolar lipids & cholesterol

Morgan et al. 1989 / Lai et al. 1995 References

a.b.c.d.

10 mL n-hexane/diethyl ether (95:5)25 mL n-hexane/diethyl ether (90:10) 15 mL n-hexane/diethyl ether (80:20) 10 mL acetone or 5 mL acetone

Chen et al. 2012, Lozada-Castro et al. 2011; Chen et al. 2010; Lee et al. 2008; Lee et al 2006; Thurner et al. 2007.

Larkeson et al. 2000 / Ubhayasekera et al. 2004 References

a.b.d.

2 mL n-hexane/diethyl ether (75:25)3 mL n-hexane/diethyl ether (60:40) 4 mL acetone or 4 mL acetone/methanol (60:40)

Ubhayasekera et al. 2012; Ubhayasekera et al. 2010; Varleyen et al. 2003.

Rose-Sallin et al. 1995 References

e. f.g.

6 mL n-hexane/ethyl acetate (95:5)10 mL n-hexane/ethyl acetate (90:10) 10 mL acetone

Cardenia et al. 2011; Boselli et al. 2010; Verardo et al. 2010; Boselli et al. 2009;

Ulberth et al. 1998 References

a. b.c. d.

f.g.

10 mL n-hexane/diethyl ether (95:5)30 mL n-hexane/diethyl ether (90:10) 10 mL n-hexane/diethyl ether (80:20) 10 mL methanol/acetone (60:20) & re-dissolved in n-hexane/ethyl acetate (90:10) 15 mL n-hexane/ethyl acetate (90:10) 10 mL acetone

Rodriguez-Carpena et al. 2012; Clariana et al. 2011a; Broncano et al. 2009; Petron et al. 2003

⁄Ulberth, Morgan and Lai methods are applied for purification of the lipid extract while the others for the enrichment of the unsaponifiable matter.

C.A. Georgiou et al. / Food Chemistry 145 (2014) 918–926 923

It is worth to mention that some authors proposed the replace-ment of cold saponification by transesterification, a method thatuses milder conditions, in order to minimize the possibility of deg-radation of vulnerable COPs by prolonged contact with alkali.Nonetheless, there is a lack of studies performed by the use of thisalternative procedure for the isolation of COPs from food matrices(Bodin & Diczfalusy, 2002; Calderón-Santiago, Peralbo-Molina, Pri-ego-Capote, Luque, & de Castro, 2012; Ubhayasekera et al., 2004).

As a conclusion, these studies underlined that the developmentof an analytical method involving saponification, is necessary to beevaluated in terms of all saponification factors including time, con-centration of KOH, temperature and sample weight. The monitor-ing of artifact generation during preparation requiresconsiderable attention in order to guarantee reliable results.

4. Solid phase extraction

The majority of published articles report the use of SPE for COPsenrichment. SPE can be applied either directly to the crude lipidextract or to the non-saponifiables for further purification (Guardi-ola et al., 2002). In regard to the polarity of different constituents ofthe lipid fraction, cholesterol esters and triacylglycerols are theleast polar, phospholipids are the most polar and cholesterol andits oxidation products are in between (Ulberth and Rossler,1998). Since SPE exploits differences in the polarity of interferingcompounds and analytes, adequate separation is achieved by step-wise elution with solvents of increasing polarity (Busch and King,2010). Thereby, enrichment of COPs can be successfully accom-plished with a suitable choice of solvents. Generally, the purifica-tion with SPE is achieved by use of the following idea; retentionof COPs on the stationary phase (usually polar) after loading thesample, removal of the interfering neutral lipids (cholesterol esterand triacylglycerols) by washing the column with an apolar solventsystem and elution of the retained cholesterol derivatives with amedium polarity solvent. Phospholipids, which are the most polar

compounds, are strongly retained on the column (Calderón-Santi-ago et al. 2012).

Usually, a chromatographic purification is preferred over sapon-ification. (Chen et al.2010). Except from being simple, low-cost andnon time-consuming, it minimizes artifact generation and/orbreakdown of COPs since the sample is not subjected to any con-tact with hot or cold alkaline solution. Another important advan-tage is the capability of removing, to a large extent, cholesterol,which can interfere with the detection and quantification of COPs(Clariana et al., 2011; Guardiola, Codony, Rafecas, & Boatella,1995). However, it should be mentioned that without previoussaponification, the total amount of COPs in a food sample can beunderestimated. This can happen because, under certain oxidationconditions, autoxidation of cholesterol fatty acid esters is fasterthan that of free cholesterol, and consequently, a significantamount of COPs may exist as fatty acid esters, which cannot bequantified (Angulo, Romera, Ramirez, & Gil 1997). Therefore,saponification is recommended in order to avoid the analytical er-ror mentioned above.

Several SPE methods have been developed for selective isolationof COPs prior to chromatographic analysis. Many types of sorbentshave been recommended with silica (Si–) and aminopropyl (NH2�)being the two predominant options, either alone or in combina-tion. On the other hand, apolar sorbents like octadecyl-modifiedsilica (ODS–) have not found widespread use (Clariana et al.,2011; Ulberth and Rossler, 1998). There are also a few cases wheresilicic acid home-made columns were constructed for the purifica-tion of COPs (Nam et al., 2001). In this part of the review, differentSPE methods are summarized with more emphasis given on thosethat are most frequently used, until nowadays.

Guardiola et al. (1995) compared four different elution systemsin order to find the one that is the most effective in regard to recov-ery. The most effective provides maximum recovery of five COPsfrom Si-SPE cartridge (7b-OH, a-CE, triol, 7-keto and 25-OH), andit simultaneously minimizes cholesterol recovery in oxysterol frac-tion (Fig. 3). Elution systems I and II suffered from low recoveries of

Fig. 4. Work-up diagram of the 3-fold SPE procedure proposed by Janoskza 2010 for the recovery of COPs from the lipid extract (CHCl3: chloroform; DE: diethyl ether; HE: n-hexane; IPA: 2-propanol; MeOH: methanol).

924 C.A. Georgiou et al. / Food Chemistry 145 (2014) 918–926

COPs, especially in the case of triol, which is the most polar COP. Inthis case, triol was not recovered at all, due to the low polarity ofthe extraction solvent. In contrast, systems III (a modified versionof Morgan & Armstrong, 1989)) and IV, provided the highest recov-eries of the different COPs due to an increase in polarity of the lastsolvent used for COPs elution (Morgan & Armstrong, 1989). Asshown in Fig. 3, the two latter sequences also applied an additionalwashing step with n-hexane/diethyl ether (80:20, v/v) prior to finalisolation. This solvent mixture was essential for the optimum elim-ination of cholesterol from oxysterol fraction. System IV was finallyconsidered as the optimum SPE method chosen because of betterprecision. However, Guardiola’s SPE-method has not found a wide-spread use. In contrast, the original elution sequence suggested byMorgan et al. (1989) is frequently employed for the purification oflipid extract. Likewise, Larkeson, Dutta, and Hansson (2000) en-riched COPs further from the non-saponifiable fraction of mincedmeat products by a Si-SPE. Nevertheless, the elution proportionswere entirely different, with a significant reduction of solvent vol-umes, in relation to those mentioned above (Table 2) (Larkesonet al., 2000; Verleyen et al., 2003).

Rose-Sallin, Huggett, Bosset, Tabacchi, and Fay (1995) describeda clean-up procedure of the unsaponifiables, which were obtainedafter direct saponification of milk powder, by using an NH2-SPE.The dried extract was dissolved in 1 mL of hexane/ethyl acetate(95:5 v/v) and was loaded into an NH2 cartridge, which was previ-ously activated with 3 mL of hexane. The residual apolar com-pounds together with cholesterol were eliminated by washingthe cartridge with the solvent sequence demonstrated in Table 2.The COPs fraction was eluted with acetone (Rose-Sallin, Huggett,Bosset, Tabacchi, & Fay, 1995). This method was widely used as avery efficient way to clean-up and extract COPs from the unsapo-nifiables of several matrices.

In a recent study performed by Calderón-Santiago et al. (2012),an NH2-SPE was also employed for COPs concentration from milkafter saponification of the lipid extract. Particularly, the obtainedunsaponifiable fraction was reconstituted in 500 lL chloroformand was applied into the cartridge, which was preconditioned bytwo consecutive washing steps with n-hexane. After the elimina-tion of apolar substances with hexane, different proportions of ace-tone, n-hexane and ethyl acetate were examined in order to

achieve the optimum elution of COPs. A mixture of n-hexane/ethylacetate (50:50, v/v) was finally proven to be the best.

In a comprehensive study, Ulberth and Rossler (1998) evaluatedthe efficiency of seven SPE methods in terms of sample capacity,purity of the extract, and recovery of COPs from spiked milk fat.The variables, in this study, were not only the solvent eluent sys-tems, but also the packing material of the cartridge. SPE clean-upprotocol using NH2 sorbent, provided efficient removal of triglycer-ides, while it was incapable of eliminating cholesterol and/or par-tial glycerides from COPs extract. On the other hand, C18 cartridgeled to greater contamination of COPs fraction. Same results withthe latter were observed when a silica cartridge was used withn-hexane/diethyl ether mixtures at ratios of 8:2 and 1:1, respec-tively. However, the use of a silica cartridge in combination withthe elution system of Morgan et al., resulted in the cleanest extract.In addition, they proposed a combination of the optimum Si-SPEfollowed by an NH2-SPE in order to reduce contaminants to a fur-ther degree. This conjugated method proved to be the most suit-able for the removal of matrix components, and, at the sametime, it provided high recovery of COPs. The main drawback wasthe low recovery of triol during NH2-SPE. The method mentionedabove was also used by Petrón et al. (2003) and Rodriguez-Carpe-na, Morcuende, Petrón, and Estevez (2012) with little modifica-tions in the sample volume (10 mg extracted lipids instead of500 mg).

Another SPE combination study was proposed by Ferioli et al.(2008). COPs were enriched from unsaponifiable material by useof two successive Si-SPE. The procedure was repeated twice in or-der to achieve a better purification of COPs-containing fractionfrom apolar materials and cholesterol. Particularly, each cartridgewas pre-equilibrated with 3 ml of n-hexane, and after fat loading,it was washed with 3 ml of n-hexane/diethyl ether (3:1, v/v) and3 ml of n-hexane/diethyl ether (3:2, v/v). The first two fractionswere discarded, and COPs were eluted with acetone/methanol(3:2, v/v).

Janoszka (2010) adapted a multistage SPE-clean up procedure,based on the method described by Regueiro and Maraschiello(1997), in order to concentrate COPs from the lipid extract (Regue-iro & Maraschiello, 1997). This unique threefold SPE procedure in-volved the isolation of sterols from the lipid fraction by using a

C.A. Georgiou et al. / Food Chemistry 145 (2014) 918–926 925

combination of SPE mega elut columns prior to final extraction ofoxysterols. COPs were further purified with a Si-SPE column(500 mg) as illustrated in Fig. 4.

5. Conclusion

In the light of the above, the sample preparation constitutes themost crucial step in COPs analysis. Even though in recent yearsnumerous techniques of extraction and purification have beenpublished, the choice between them remains a problem. The gen-eral conclusion of the current review is that all sample preparationparameters (i.e., type of solvents for the lipid extraction, the time ofsaponification and the elution systems or type of sorbents of SPE)must be evaluated prior to application. This aspect is essential inorder to achieve an efficient recovery of COPs from the food matrixand to provide the most reliable results. The existence of severalmodified versions demonstrated that even minor changes, duringthe procedure, can influence the final quantification of COPs.Emphasis should also be given on artifact formation. This is a veryimportant parameter that is usually omitted, leading to inaccuratedata. In addition, interlaboratory comparisons of evaluation andquantitation data can be performed in order to improve the meth-odologies, eliminate the analytical errors, and consequently, devel-op a reliable and certified methodology for the analysis of COPs infood samples.

References

Angulo, A. J., Romera, J. M., Ramirez, M., & Gil, A. (1997). Determination ofCholesterol Oxides in Dairy Products. Effect f Storage Conditions. Journal ofAgricultural and Food Chemistry, 45, 4318–4323.

Baggio, S. R., & Bragagnolo, N. (2006). Cholesterol oxide, cholesterol, total lipid andfatty acid contents in processed meat products during storage. LWT, 39,513–520.

Baggio, S. R., Miguel, A. M. R., & Bragagnolo, N. (2005). Simultaneous determinationof cholesterol oxides, cholesterol and fatty acids in processed turkey meatproducts. Food Chemistry, 89, 475–484.

Bielska, A. A., Schlesinger, P., Covey, D. F., & Ory1, D. S. (2012). Oxysterols as non-genomic regulators of cholesterol homeostasis. Trends in Endocrinology andMetabolism, 23, 99–106.

Bligh, E. G., & Dyer, W. J. (1959). A rapid method of total lipid extraction andpurification. Canadian Journal of Biochemistry and Physiology, 37, 911–917.

Bodin, K., & Diczfalusy, U. (2002). Analysis of cholesterol oxidation products inplasma, tissues and food. European Journal of Lipid Science and Technology, 104,435–439.

Bonoli, M., Caboni, M. F., Rodriguez-Estrada, M. T., & Lercker, G. (2007). Effect offeeding fat sources on the quality and composition of lipids of precooked ready-to-eat fried chicken patties. Food Chemistry, 101, 1327–1337.

Boselli, E., Caboni, M. F., Rodriguez-Estrada, M. T., Toschi, T. G., Daniel, M., & Lercker,G. (2005). Photoxidation of cholesterol and lipids of turkey meat during storageunder commercial retail conditions. Food Chemistry, 91, 705–713.

Boselli, E., Rodriguez-Estrada, M. T., Fedrizzi, G., & Caboni, M. F. (2009). Cholesterolphotosensitized oxidation of beef meat under standard and modifiedatmosphere at retail conditions. Meat Science, 81, 224–229.

Boselli, E., Velazco, V., Caboni, M. F., & Lercker, G. (2001). Pressurized liquidextraction of lipids for the determination of oxysterols in egg-containing food.Journal of Chromatography A, 917, 239–244.

Bosselli, E., Rodriguez-Estrada, M. T., Ferioli, F., Caboni, M. F., & Lercker, G. (2010).Cholesterol photosensitized oxidation of horse meat slices stored underdifferent packaging films. Meat Science, 85, 500–505.

Broncano, J. M., Petrón, M. J., Parra, V., & Timón, M. L. (2009). Effect of differentcooking methods on lipid oxidation and formation of free cholesterol oxidationproducts (COPs) in Latissimus dorsi muscle of Iberian pigs. Meat Science, 83,431–437.

Brown, A. J., & Jessup, W. (1999). Oxysterols and atherosclerosis. Atherosclerosis, 142,1–28.

Brown, A. J., & Jessup, W. (2009). Oxysterols: Sources, cellular storage andmetabolism, and new insights into their roles in cholesterol homeostasis.Molecular Aspects of Medicine, 30, 111–122.

Busch, T. P., & King, A. J. (2009). Artifact generation and monitoring in analysis ofcholesterol oxide products. Analytical Biochemistry, 388, 1–14.

Busch, T. P., & King, A. J. (2010). Stability of cholesterol, 7-ketocholesterol and b-sitosterol during saponification: Ramifications for artifact monitoring of steroloxide products. Journal of the American Oil Chemists Society, 87, 955–962.

Calderón-Santiago, M., Peralbo-Molina, Á., Priego-Capote, F., Luque, Dolores, & deCastro, M. (2012). Cholesterol oxidation products in milk: Processing formationand determination. European Journal of Lipids Science and Technology, 114,687–694.

Cardenia, V., Rodriguez-Estrada, M. T., Baldacci, E., Savioli, S., & Lercker, G. (2012).Analysis of cholesterol oxidation products by fast gas chromatography/massspectrometry. Journal of Separation Science, 35, 424–430.

Cardenia, V., Rodriguez-Estrada, M. T., Cumella, F., Sardi, L., Casa, C. D., & Lercker, G.(2011a). Oxidative stability of pork meat lipids as related to high-oleicsunflower oil and vitamin E diet supplementation and storage conditions.Meat Science, 88, 271–279.

Chen, Y. C., Chien, J. T., Inbaraj, B. S., & Chen, B. H. (2012). Formation and inhibitionof cholesterol oxidation products during marinating of pig feet. Journal ofAgricultural and Food Chemistry, 60, 173–179.

Chen, L. J., Lu, Y. F., Chien, J. T., & Chen, B. H. (2010). Formation and inhibition ofcholesterol oxidation products in tea-leaf eggs during marinating. Journal ofAgricultural and Food Chemistry, 58, 10467–10474.

Christie, W. W. (1993). Preparation of lipid extracts from tissues. Advances in LipidMethodology, 2, 195–213.

Clariana, M., Díaz, I., Sárraga, C., & García-Regueiro, J. (2011). Comparison of thedetermination of eight cholesterol oxides in dry–cured shoulder by GC–FID,GC–MS, and GC tandem mass spectrometry. Food Analytical Methods, 4,465–474.

Clariana, M., & García-Regueiro, J. A. (2011). Effect of high pressure processing oncholesterol oxidation products in vacuum packaged sliced dry-cured ham. Foodand Chemical Toxicology, 49, 1468–1471.

Derewiaka, D., & Obiedzinski, M. (2010). Cholesterol oxides content in selectedanimal products determined by GC-MS. European Journal of Lipid Science andTechnology, 112, 1130–1137.

Dionisi, F., Golay, P. A., Aeschlimann, J. M., & Fay, L. B. (1998). Determination ofcholesterol oxidation products in milk powders: methods comparison andvalidation. Journal of Agricultural and Food Chemistry, 46, 2227–2233.

Du, M., Nam, K. C., & Ahn, D. U. (2001). Cholesterol and lipid oxidation products incooked meat as affected by raw-meat packaging and irradiation and by cooked-meat packaging and storage time. Journal of Food Science, 66, 1396–1401.

Echarte, M., Ansorena, D., & Astiasarán, I. (2003). Consequences of microwaveheating and frying on the lipid fraction of chicken and beef patties. Journal ofAgricultural and Food Chemistry, 51, 5941–5945.

Farese, R. V., & Walther, T. C. (2009). Lipid Droplets Finally Get a Little R-E-S-P-E-C-T. Cell, 139, 855–860.

Ferioli, F., Caboni, M. F., & Dutta, P. C. (2008). Evaluation of cholesterol and lipidoxidation in raw and cooked minced beef stored under oxygen-enrichedatmosphere. Meat Science, 80, 681–685.

Ferioli, F., Dutta, P. C., & Caboni, M. F. (2010). Cholesterol and lipid oxidation in rawand pan-fried minced beef stored under aerobic packaging. Journal of the Scienceof Food and Agriculture, 90, 1050–1055.

Folch, J., Lees, M., & Stanley, G. H. S. (1957). A simple method for the isolation andpurification of total lipides from animal tissues. Journal of Biological Chemistry,226, 497–509.

Georgiou, C. A., & Kapnissi-Christodoulou, C. P. (2013). Qualitative and quantitativedetermination of COPs in Cypriot meat samples using HPLC. Determination ofthe most effective sample preparation procedure. Journal of ChromatographicScience, 51, 286–291.

Gill, S., Chow, R., & Brown, A. J. (2008). Sterol regulators of cholesterol homeostasisand beyond: The oxysterol hypothesis revisited and revised. Progress in LipidResearch, 47, 391–404.

Grau, A., Cobony, R., Grimpa, S., Baucells, M. D., & Guardiola, F. (2001). Cholesteroloxidation in frozen dark chicken meat: influence of dietary fat source, and a-tocopherol and ascorbic acid supplementation. Meat Science, 57, 197–208.

Guardiola, F., Codony, R., Rafecas, M., & Boatella, J. (1995). Comparison of threemethods for the determination of oxysterols in spray-dried egg. Journal ofChromatography A, 705, 289–304.

Guardiola, F., Dutta, P. C., Codony, R., & Savage, G. P. (2002). Cholesterol andPhytosterol Oxidation Products Analysis, Occurrence, and Biological Effects.AOCS Press, (Chapter 3).

Hara, A., & Radin, N. S. (1978). Lipid extraction of tissues with a low toxicity solvent.Analytical Biochemistry, 90, 420–426.

Hur, S. J., Park, G. B., & Joo, S. T. (2007). Formation of cholesterol oxidation products(COPs) in animal products. Food Control, 18, 939–947.

Iverson, S. J., Lang, S. L. C., & Cooper, M. H. (2001). Comparison of the Bligh and Dyerand Folch Methods for Total Lipid Determination in a Broad Range of MarineTissue. Lipids, 36, 1283–1287.

Janoszka, B. (2010). 7-ketocholesterol and 7-hydroxycholesterol in pork meat andits gravy thermally treated without additives and in the presence of onion andgarlic. Meat Science, 86, 976–984.

Jensen, M. Ø., & Mouritsen, O. G. (2004a). Lipids do influence protein function - thehydrophobic matching hypothesis revisited. Biochimica et Biophysica Acta, 1666,205–226.

Jusakul, A., Yongvanit, P., Loilome, W., Namwat, N., & Kuver, R. (2011). Mechanismsof oxysterol-induced carcinogenesis. Lipids in Health and Disease, 2011(10),44–51.

Larkeson, B., Dutta, P. C., & Hansson, I. (2000). Effects of frying and storage oncholesterol oxidation in minced meat products. Journal of the American OilChemists Society, 77, 675–680.

926 C.A. Georgiou et al. / Food Chemistry 145 (2014) 918–926

Lee, H.-W., Chien, J.-T., & Chen, B.-T. (2006). Formation of cholesterol oxidationproducts in marinated foods during heating. Journal of Agricultural and FoodChemistry, 54, 4873–4879.

Lee, H. W., Chien, J. T., & Chen, B. H. (2008). Inhibition of cholesterol oxidation inmarinated foods as affected by antioxidants during heating. Food Chemistry, 108,234–244.

Leonarduzzi, G., Sottero, B., & Poli, G. (2002). Oxidized products of cholesterol:dietary and metabolic origin, and proatherosclerotic effects (review). Journal ofNutritional Biochemistry, 13, 700–710.

Lordan, S., Mackrill, J. J., & O’Brien, N. M. (2009). Oxysterols and mechanisms ofapoptotic signaling: implications in the pathology of degenerative diseases.Journal of Nutritional Biochemistry, 20, 321–336.

Lozada-Castro, J. J., Gil-Diaz, M., Santos-Delgado, M. J., Rubio-Barroso, S., & Polo-Diez, L. M. (2011). Effect of electron-beam irradiation on cholesterol oxideformation in different ready-to-eat foods. Innovative Food Science and EmergingTechnologies, 12, 519–525.

Manini, P., Andreoli, R., Careri, M., Elviri, L., & Musci, M. (1998). Atmosphericpressure chemical ionization liquid chromatography/mass spectrometry incholesterol oxide determination and characterization. Rapid Communications inMass Spectrometry, 12, 883–889.

Mariutti, L. R. B., Nogueira, G. C., & Bragagnolo, N. (2008). Optimization andvalidation of analytical conditions for cholesterol oxides extraction in chickenmeat using response surface methodology. Journal of Agricultural and FoodChemistry, 56, 2913–2918.

Maxwell, R. J., Mondimore, D., & Tobias, J. (1986). Rapid method for the quantitativeextraction and simultaneous class separation of milk lipids. Journal of DairyScience, 69, 321–325.

Mazalli, M. R., & Bragagnolo, N. (2007). Effect of storage on cholesterol oxideformation and fatty acid alterations in egg powder. Journal of Agricultural andFood Chemistry, 55, 2743–2748.

Mazalli, M. R., & Bragagnolo, N. (2009). Increase of cholesterol oxidation anddecrease of PUFA as a result of thermal processing and storage in eggs enrichedwith n-3 fatty acids. Journal of Agricultural and Food Chemistry, 57, 5028–5034.

Mazalli, M. R., Sawaya, A. C. H. F., Eberlin, M. N., & Bragagnolo, N. (2006). HPLCmethod for quantification and characterization of cholesterol and its oxidationproducts in eggs. Lipids, 41, 615–622.

Morales-Aizpurua, I. C., & Tenuta-Filho, A. (2005). Oxidation of cholesterol inmayonnaise during storage. Food Chemistry, 89, 611–615.

Morgana, J. N., & Armstrong, D. J. (1989). Wide-bore capillary gas chromatographicmethod for quantification of cholesterol oxidation products in egg yolk powder.Journal of Food Science, 54, 427–430.

Nam, K. C., Du, M., Jo, C., & Ahn, D. U. (2001). Cholesterol oxidation products inirradiated raw meat with different packaging and storage time. Meat Science, 58,431–435.

Obara, A., Obiedzinski, M., & Kołczak, T. (2006). The effect of water activity oncholesterol oxidation in spray- and freeze-dried egg powders. Food Chemistry,95, 173–179.

Olkkonen, V. M., Béaslas, O., & Nissilä, E. (2012). Oxysterols and Their CellularEffectors. Biomolecules, 2, 76–103.

Olsen, B. N., Schlesinger, P. H., & Baker, N. A. (2009). Perturbations of membranestructure by cholesterol and cholesterol derivatives are determined by sterolorientation. Journal of American Chemical Society, 131, 4854–4865.

Olsen, B. N., Schlesinger, P. H., Ory, D. S., & Baker, N. A. (2012). Side-chain oxysterols:From cells to membranes to molecules. Biochimica et Biophysica Acta, 1818,330–336.

Otaegui-Arrazola, A., Menendez-Carreño, M., Ansorena, D., & Astiasarán, I. (2010).Oxysterols: A world to explore. Food and Chemical Toxicology, 48, 3289–3303.

Park, W. T., Guardiola, F., Park, S. H., & Addis, P. B. (1996). Kinetic evaluation of 3b-hydroxycholest-5-en-7-one (7-ketocholesterol) stability during saponification.Journal of the American Oil Chemists Society, 73, 623–629.

Petrón, M. J., Garcia-Regueiro, J. A., Martin, L., Muriel, E., & Antequera, T. (2003).Identification and quantification of cholesterol and cholesterol oxidationproducts in different types of Iberian hams. Journal of Agricultural and FoodChemistry, 51, 5786–5791.

Pignoli, G., Rodriguez-Estrada, M. T., Mandrioli, M., Barbanti, L., Rizzi, L., & Lercker,G. (2009). Effects of different rearing and feeding systems on lipid oxidation andantioxidant capacity of freeze-dried egg yolks. Journal of Agricultural and FoodChemistry, 57, 11517–11527.

Poli, G., Sottero, B., Gargiulo, S., & Leonarduzzi, G. (2009). Cholesterol oxidationproducts in the vascular remodeling due to atherosclerosis. Molecular Aspects ofMedicine, 30, 180–189.

Raith, K., Brenner, C., Farwanah, H., Müller, G., Eder, K., & Neubert, R. H. H. (2005). Anew LC/APCI-MS method for the determination of cholesterol oxidationproducts in food. Journal of Chromatography A, 1067, 207–211.

Regueiro, J. A. G., & Maraschiello, C. (1997). Procedure for the determination of eightcholesterol oxides in poultry meat using on-column and solvent ventingcapillary gas chromatography. Journal of Chromatography A, 764, 279–293.

Reue, K. (2011). A Thematic Review Series: Lipid droplet storage and metabolism:from yeast to man. Journal of Lipid Research, 52, 1865–1868.

Rodriguez-Carpena, J.-G., Morcuende, D., Petrón, M. J., & Estevez, M. (2012).Inhibition of cholesterol oxidation products (COPs) formation in emulsifiedporcine patties by phenolic-rich avocado (Persea americana Mill.) extracts.Journal of Agricultural and Food Chemistry, 60, 2224–2230.

Rose-Sallin, C., Huggett, A. C., Bosset, J. O., Tabacchi, R., & Fay, L. B. (1995).Quantification of cholesterol oxidation products in milk powders using [2H7]cholesterol to monitor cholesterol autoxidation artifacts. Journal of Agriculturaland Food Chemistry, 43, 935–941.

Ruiz-Gutierrez, V., & Perez-Camino, M. C. (2000). Update on solid-phase extractionfor the analysis of lipid classes and related compounds. Journal ofChromatography A, 885, 321–341.

Saldanha, T., Benassi, M. T., & Bragagnolo, N. (2008). Fatty acid contents evolutionand cholesterol oxides formation in Brazilian sardines (Sardinella brasiliensis) asa result of frozen storage followed by grilling. LWT, 41, 1301–1309.

Saldanha, T., & Bragagnolo, N. (2007). Cholesterol oxidation is increased and PUFAdecreased by frozen storage and grilling of atlantic hake fillets (Merlucciushubbsi). Lipids, 42, 671–678.

Saldanha, T., Sawaya, A. C. H. F., Eberlin, M. N., & Bragagnolo, N. (2006). HPLCseparation and determination of 12 cholesterol oxidation products in fish:Comparative study of RI, UV, and APCI-MS detectors. Journal of Agricultural andFood Chemistry, 54, 4107–4113.

Sampaio, G. R., Bastos, D. H. M., Soares, R. A. M., Queiroz, Y. S., & Torres, E. A. F. S.(2006). Fatty acids and cholesterol oxidation in salted and dried shrimp. FoodChemistry, 95, 344–351.

Savage, G. P., Dutta, P. C., & Rodriguez-Estrada, M. T. (2002). Cholesterol oxides:their occurrence and methods to prevent their generation in foods. Asia PasificJournal of Clinical Nutrition, 11, 72–78.

Sieber, R. (2005). Oxidised cholesterol in milk and dairy products. InternationalDairy Journal, 15, 191–206.

Soto-Rodriguez, I., Campillo-Velazquez, J. P., Ortega-Martinez, J., Rodriguez-Estrada,M. T., Lercker, G., & Garcia, S. H. (2008). Cholesterol oxidation in traditionalMexican dried and deep-fried food products. Journal of Food Composition andAnalysis, 21, 489–495.

Tai, C.-Y., Chen, Y. C., & Chen, B. H. (1999). Analysis, formation and inhibition ofcholesterol oxidation products in foods: An overview (Part I). Journal of Food andDrug Analysis, 7, 243–257.

Ubhayasekera, J. K. A. S., Jayasinghe, P., Ekanayake, S., & Paresh, C. D. (2012). Highcholesterol oxidation in pickled mackerel (Rastrelliger kanagurta) from SriLanka. European Journal of Lipid Science and Technology, 114, 695–700.

Ubhayasekera, S. J. K. A., Tres, A., Cobony, R., & Dutta, P. C. (2010). Effects of differentlevels of trans fatty acids and oxidised lipids in diet on cholesterol andcholesterol oxidation products formation in rabbit. Food Chemistry, 121,1198–1202.

Ubhayasekera, S. J. K. A., Verleyen, T., & Dutta, P. C. (2004). Evaluation of GC and GC-MS methods for the analysis of cholesterol oxidation products. Food Chemistry,84, 149–157.

Ulberth, F., & Rossler, D. (1998). Comparison of solid phase extraction methods forthe cleanup of cholesterol oxidation products. Journal of Agricultural and FoodChemistry, 46, 2634–2637.

Verardo, V., Pasini, F., Iafelice, G., Messia, M. C., Marconi, E., & Caboni, M. F. (2010).Influence of storage conditions on cholesterol oxidation in dried egg pasta.Journal of Agricultural and Food Chemistry, 58, 3586–3590.

Verleyen, T., Dutta, P. C., Verhe, R., Dewettinck, K., Huyghebaert, A., & Greyt, D. W.(2003). Cholesterol oxidation in tallow during processing. Food Chemistry, 83,185–188.

Vicente, S. J. V., Sampaio, G. R., Ferrari, C. K. B., & Torres, E. A. F. S. (2012). Oxidationof cholesterol in foods and its importance for human health. Food ReviewsInternational, 28, 47–70.

Vicente, S. J. V., & Torres, E. A. F. S. (2007). Formation of four cholesterol oxidationproducts and loss of free lipids, cholesterol and water in beef hamburgers as afunction of thermal processing. Food Control, 18, 63–68.

Yen, T. Y., Inbaraj, B. S., Chien, L. T., & Chen, B. H. (2010). Gas chromatography-massspectrometry determination of conjugated linoleic acids and cholesterol oxidesand their stability in a model system. Analytical Biochemistry, 400, 130–138.