Phytosterols, phytostanols, and their conjugates in foods: structural diversity, quantitative...

Review

Phytosterols, phytostanols, and their conjugates in foods:structural diversity, quantitative analysis, and

health-promoting uses§

Robert A. Moreaua,*, Bruce D. Whitakerb, Kevin B. Hicksa

aCrop Conversion Science and Technology Research Unit, Eastern Regional Research Center, United States Departmentof Agriculture, Agricultural Research Service, 600 East Mermaid Lane, Wyndmoor, PA 19038, USA

bProduce Quality and Safety Laboratory, Beltsville Agricultural Research Center, United States Department of Agriculture,Agricultural Research Service, 10300 Baltimore Avenue, Beltsville, MD 20705, USA

Received 1 February 2002; received in revised form 15 March 2002; accepted 22 March 2002

Abstract

Phytosterols (plant sterols) are triterpenes that are important structural components of plant membranes,and free phytosterols serve to stabilize phospholipid bilayers in plant cell membranes just as cholesteroldoes in animal cell membranes. Most phytosterols contain 28 or 29 carbons and one or two carbon–carbondouble bonds, typically one in the sterol nucleus and sometimes a second in the alkyl side chain. Phytos-tanols are a fully-saturated subgroup of phytosterols (contain no double bonds). Phytostanols occur intrace levels in many plant species and they occur in high levels in tissues of only in a few cereal species.Phytosterols can be converted to phytostanols by chemical hydrogenation. More than 200 different typesof phytosterols have been reported in plant species. In addition to the free form, phytosterols occur as fourtypes of ‘‘conjugates,’’ in which the 3b-OH group is esterified to a fatty acid or a hydroxycinnamic acid, orglycosylated with a hexose (usually glucose) or a 6-fatty-acyl hexose. The most popular methods for phy-tosterol analysis involve hydrolysis of the esters (and sometimes the glycosides) and capillary GLC of thetotal phytosterols, either in the free form or as TMS or acetylated derivatives. Several alternative methodshave been reported for analysis of free phytosterols and intact phytosteryl conjugates. Phytosterols andphytostanols have received much attention in the last five years because of their cholesterol-loweringproperties. Early phytosterol-enriched products contained free phytosterols and relatively large dosageswere required to significantly lower serum cholesterol. In the last several years two spreads, one containingphytostanyl fatty-acid esters and the other phytosteryl fatty-acid esters, have been commercialized and

0163-7827/02/$ - see front matter Published by Elsevier Science Ltd.

PI I : S0163-7827(02 )00006-1

Progress in Lipid Research 41 (2002) 457–500

www.elsevier.com/locate/plipres

§ Mention of a brand or firm name does not constitute an endorsement by the US Department of Agriculture aboveothers of a similar nature not mentioned.* Corresponding author.

E-mail address: [email protected] (R.A. Moreau).

were shown to significantly lower serum cholesterol at dosages of 1–3 g per day. The popularity ofthese products has caused the medical and biochemical community to focus much attention on phytosterolsand consequently research activity on phytosterols has increased dramatically. Published by ElsevierScience Ltd.

Contents

1. Introduction and a primer on phytosterol nomenclature....................................................................................... 459

2. Structural diversity and phylogenetic distribution of phytosterols ........................................................................ 4652.1. Occurrence and metabolism of cholesterol in plants ..................................................................................... 4652.2. Distribution and diversity of major C-24 alkyl phytosterols......................................................................... 466

2.3. Free and conjugated phytosterols in fruits and vegetables............................................................................ 4672.4. Unique phytosterols and phytosterol conjugates in cereals........................................................................... 4702.5. Function of 24-alkyl phytosterols and their conjugates ................................................................................ 470

2.6. Steroidal saponins ......................................................................................................................................... 4712.7. Steroidal glycoalkaloids................................................................................................................................. 4722.8. Phytoecdysteroids and brassinosteroids ........................................................................................................ 474

3. Methods for the Quantitative Analysis of Phytosterols and Phytostanols............................................................. 4763.1. Extraction and fractionation of phytosterols ................................................................................................ 4763.2. Methods for the separation of intact phytosterol classes .............................................................................. 477

3.3. Methods for the separation of molecular species of phytosterol conjugates ................................................. 4783.4. Methods for hydrolysis of conjugates and separation of free phytosterols ................................................... 4793.5. Mass spectrometry and NMR of phytosterols .............................................................................................. 481

3.6. Enzymatic assays of phytosterols .................................................................................................................. 482

4. Health-promoting effects of phytosterols, phytostanols, and their esters .............................................................. 4834.1. Historical perspectives, changing dogma, and critical questions................................................................... 483

4.2. Active forms and mechanism of action of steryl and stanyl esters................................................................ 4854.3. Recent clinical studies on phytostanyl esters................................................................................................. 4854.4. Recent clinical studies on phytosteryl esters.................................................................................................. 486

4.5. Recent clinical studies on free phytosterols and phytostanols....................................................................... 4864.6. Relative LDL-C lowering efficacy: sterols vs stanols & esters vs free forms ................................................. 4874.7. Health claims................................................................................................................................................. 488

4.8. Reduction in the risk of coronary heart disease ............................................................................................ 4884.9. Toxicology, anticancer properties and potential benefits .............................................................................. 4884.10. Effects on absorption of fat soluble vitamins and antioxidants .................................................................... 489

4.11. Dosage levels and frequency.......................................................................................................................... 4904.12. Phytosterol/phytostanol products: past, present, and in development .......................................................... 491

5. Conclusions ............................................................................................................................................................ 494

Acknowledgements...................................................................................................................................................... 494

References ................................................................................................................................................................... 495

458 R.A. Moreau et al. / Progress in Lipid Research 41 (2002) 457–500

1. Introduction and a primer on phytosterol nomenclature

Phytosterols (plant sterols) are members of the ‘‘triterpene’’ family of natural products, whichincludes more than 100 different phytosterols and more than 4000 other types of triterpenes [1,2].Cholesterol is the predominant sterol in animals, wherein free cholesterol serves to stabilize cellmembranes and cholesteryl fatty-acid esters are a storage/transport form, usually found associatedwith triacylglycerols [3]. Plant membranes contain little or no cholesterol and instead contain severaltypes of phytosterols that are similar in structure to cholesterol but include a methyl or ethylgroup at C-24. In general, phytosterols are also thought to stabilize plant membranes, with anincrease in the sterol/phospholipid ratio leading to membrane rigidification [4]. However, indivi-dual phytosterols differ in their effect on membranes stability. Stigmasterol has been reported tohave a disordering effect on membranes [5] and the molar ratio of stigmasterol to other phytos-terols in the plasma membrane increases during senescence [6]. All triterpenes are synthesized via apathway that starts with reduction of HMG-CoA (six carbons) to mevalonate (five carbons). Sixmevalonate units are then assembled into two farnesyl diphosphate molecules, which are combinedto make squalene (30 carbons or ‘‘three terpenes’’). Enzymatic ring closure steps then formcycloartenol (also 30 carbons), and additional enzymatic reactions form common plant triterpenessuch as phytosterols, triterpene alcohols, and brassinosteroids (Fig. 1).There have been several excellent reviews on phytosterols, most notably a review by Goad [1],

which focused on phytosterol chemistry and analytical methods, and one by Piironen and col-leagues [3], which was a very comprehensive review of biological, chemical, and nutrition aspectsof phytosterols. The purpose of this review is not to be a comprehensive treatise on all of phy-tosterol chemistry, structure and function, but to focus on three areas of phytosterol research thatmay be of interest to scientists largely unacquainted with the field. The three topics are as follows:(A) the occurrence of various phytosterols in plants and a discussion of the nomenclature, (B)

Nomenclature

ASG acylated steryl glycosideDGDG digalactosyldiacylglycerolFFA free (non-esterified) fatty acidsFS free sterolFSE ferulate steryl esterHDL-C high density lipoprotein serum cholesterolHSE hydroxycinnamate steryl esterLDL-C low density lipoprotein serum cholesterolLyso-PC lysophosphatidylcholineSE steryl fatty-acid esterPE phosphatidylethanolaminePI phosphatidylinositolPC phosphatidylcholineSG steryl glycosideTAG triacylglycerol

R.A. Moreau et al. / Progress in Lipid Research 41 (2002) 457–500 459

modern analytical tools used to quantify and identify phytosterols and a comparison of the var-ious analytical methods, and (C) recent advances in the health-promoting and nutraceuticalaspects of phytosterols.Phytosterol nomenclature is confusing because international attempts at standardization have

been only partially adopted. The two main nomenclatures (Fig. 2) currently utilized follow theIUPAC-IUB recommendations of 1976 and 1989 [1]. Our approach in this review will be to referto phytosterols by their common (trivial) names and in the latter part of this section we will listthe common phytosterols, alternative names, systematic names, molecular masses, and CAS(Chemical Abstracts Service) registry numbers.A convenient way to describe and catalog phytosterols is to divide them into three groups based

on the number of methyl groups on carbon-4, two (4-dimethyl), one (4-monomethyl), or none(4-desmethyl). 4-Dimethylsterols and 4a-monomethylsterols are metabolic intermediates in thebiosynthetic pathway leading to end-product, 4-desmethyl phytosterols, but they are usuallypresent at low levels in most plant tissues. Cycloartenol and cycloartanol are examples of4-dimethylsterols, and gramisterol is an example of a 4a-monomethylsterol (Fig. 3).

Cycloartanol, also called 9,19-cyclo-5a, 9b-lanostan-3b-ol or cycloartan-3b-ol, C30H52O, MW428.74, CAS # 4657-58-3. Found in rice bran oil and the rhizomes of Polypodium vulgare.

Gramisterol, also called 24-methylenelophenol, C29H49O, MW 412.69, CAS# 1176-52-9.

Fig. 1. Biosynthesis of phytosterols and other triterpenes.


4-Desmethylsterols include the 27-carbon sterol cholesterol (Fig. 4) (ubiquitous and pre-dominant in animals, but also generally present in plants at low levels) and all of the common28-carbon (Fig. 5) and 29-carbon (Fig. 6) phytosterols, which are typically major membranestructural components in plant cells. Most 4-desmethyl phytosterols have a double bond betweencarbons 5 and 6 of the ring system and are thus called �5 phytosterols. However, another groupof common desmethylsterols that are abundant in plants of certain families have a double bond

Fig. 2. Nomenclature of phytosterols (example: sitosterol=stigmast-5-en-3b-ol=24R-ethylcholest-5-en-3b-ol).

Fig. 3. 4-Monomethyl and 4,4-dimethylphytosterols.


between carbons 7 and 8 instead of 5 and 6, and are hence referred to as �7 phytosterols. Both �5

and �7 desmethylsterols can include a second double bond in the alkyl side chain, most fre-quently between carbons 22 and 23 or carbons 24 and 28 (carbons 24 and 241 in the 1989 IUPACnomenclature). The common 29-carbon desmethylsterol stigmasterol (Fig. 6), which includesboth C5,6 and (trans) C22,23 double bonds, is, for example, designated as �5,22E.For the C28 and C29 phytosterols the introduction of a methyl or ethyl group at C24 renders

this position chiral and thus two epimers are possible. The nomenclature of the configuration of

Fig. 4. C27 4-desmethyl phytosterols.

Fig. 5. C28 4-desmethyl phytosterols. Note that some of these compound are 24a (solid wedge) and some are 24b(dashed wedge). Catalytic hydrogenation (a method currently used in the production of commercial stanyl ester pro-ducts) of the unsaturated C28 phytosterols can yield two 24-methyl epimers, campestanol (24a=24R) or ergostanol(24b=24S), depending on the phytosterol composition of the original material.


the C24 methyl, C24 ethyl, or C24 ethylidene groups on the C28 and C29 phytosterols requiressome explanation. For the seven common C28 phytosterols listed in Fig. 5, three (campesterol,epibrassicasterol, and campestanol) are 24a epimers (with the methyl group indicated as a ‘‘solidwedge’’), and the other five phytosterols are 24b epimers (with the methyl group indicated as a‘‘dashed wedge’’). The 24-methyl epimers are also designated 24R and 24S, which are equivalentto 24a and 24b, respectively, unless there is a double bond at C22,23, in which case the chirality isreversed (24R=24b and 24S=24a). With the C29 phytosterols, the good news is that almost allphytosterols are 24a-ethyl epimers. Unfortunately, three of the common C29 phytosterols have adouble bond at C24,28 (C24,240) and the resulting ethylidene group can either be cis or trans. TheC24 ethylidene in fucosterol is the trans isomer and is designated as a 24E, whereas the C24 eth-ylidene in �5-avenasterol and �5-avenasterol is the cis isomer and is designated as 24Z. For-tunately, the C22,23 double bond in common phytosterols (brassicasterol, epibrassicasterol,stigmasterol, and 7-stigmasterol (spinasterol) only occur as 22E.

27-Carbon 4-desmethylsterols (Fig. 4).

Cholesterol, cholest-5-en-3b-ol, C27H46O, mol. wt. 386.65, CAS# 57-88-5. The common sterolin animal tissues. Also occurs in the date palm, Phoenix dactylifera, and in many marine redalgae (Rhodophyceae).

Desmosterol, also called cholesta-5,24-dien-3b-ol, 24-dehydrocholesterol, C27H44O, MW384.63, CAS# 313-04-02.

Fig. 6. C29 4-desmethyl phytosterols. Note that catalytic hydrogenation (a method currently used in the production of

commercial stanyl ester products) of unsaturated C29 phystosterols can yield stigmastanol and/or its 24 b epimer,depending on the phytosterol composition of the original material.


Lathosterol, also called 5a-cholest-7-en-3b-ol, C27H46O, mol. wt. 386.65. Lathosterol is the C27precursor to phytoecdysteroids in spinach (Spinacia oleracea).

Cholesta-5,7-dien-3�-ol, also called 7-dehydrocholesterol, �7-cholesterol, and Provitamin D3,C27H44O, MW 384.63, CAS# 434-16-2.

28-Carbon 4-desmethylsterols (Fig. 5)

Campestanol (saturated) (24a=24R), also called (24R)-24-methylcholestan-3b-ol or (24R)-ergostan-3b-ol, C28H50O, MW 402.70, CAS # 474–60–2. Occurs naturally in corn fiber oil,almost exclusively as a ferulate or p-coumarate esters. Generated by catalytic hydrogenation ofcampesterol or epibrassicasterol.

Ergostanol (saturated) (24b=24S), also called (24S)-24-methylcholestan-3b-ol, C28H50O, MW402.70. Not reported naturally but can be generated by catalytic hydrogenation of brassicas-terol, 22-dihydrobrassicasterol, or ergosterol.

Campesterol (�5) (24a=24R), also called (24R)-24-methylcholest-5-en-3b-ol, campest-5-en-3b-ol, or �5-24a-methyl-cholesten-3b-ol, (24R)-ergost-5-en-3b-ol, C28H48O, mol. Wt. 400.68, CAS# 474-62-4. Widespread occurrence in plants.

22-Dihydrobrassicasterol (�5) (24b=24S), also called ergost-5-en-3b-ol and 24-epicampesterol,C28H48O, mol. Wt. 400.68, CAS # 4651-51-8.

Brassicasterol (�5,22E) (24b=24R), also called (22E)-ergosta-5,22-dien-3b-ol, C28H46O, MW398.66, CAS # 474-67-9. Found in rapeseed oil from Brassica napus.

Epibrassicasterol (�5,22E) (24a=24S), also called (22E)-(24S)-24-methylcholesta-5,22-dien-3b-ol or (22E)-campesta-5,22-dien-3b-ol, C28H46O, MW 398.66, CAS # 17472-78-5.

Ergosterol (�5,7,22E) (24b=24R), also called (22E)-ergosta-5,7,22-trien-3b-ol, C28H44O, MW396.54, CAS # 57-87-4. Occurs in yeasts and many other fungi.

29-Carbon 4-desmethylsterols (Fig. 6)

Sitostanol (saturated) (24a=24R), also called stigmastanol, stigmastan-3b-ol, 24a-ethylchol-estan-3b-ol, C29H52O, MW 416.40, CAS # 19466-47-8. Occurs in corn fiber oil, almost exclu-sively as a ferulate ester or p-coumarate esters.

Sitosterol (�5) (24a=24R), also called b-sitosterol, stigmast-5-en-3b-ol, 24a-ethylcholest-5-en-3b-ol, C29H50O, MW 414.71, CAS # 83-46-5. Widespread occurrence in plants.

D 7-Stigmastenol (7-Stigmastenol) (�7) (24a=24S), also called, 22-dihydrospinasterol, stig-masta-7-en-3b-ol, C29H50O, MW 414.71, CAS # 521-03-9.


Stigmasterol (�5,22E) (24a=24S), also called (22E)-stigmasta-5,22-dien-3b-ol or 24a-ethylcho-lesta-5,22E-dien-3b-ol, C29H48O, MW 412.69, CAS # 83-48-7. Widespread occurrence in plants.

Fucosterol (�5,24E), also called [24(28)E]-stigmasta-5,24(28)-dien-3b-ol, [24(240)E]-stigmasta-5,24(240)-dien-3b-ol, or 24E-ethylidenecholesta-5,24(28)-dien-3b-ol, C29H48O, MW 412.69,CAS # 17605-67-3. Found in coconut pollen, Cocos nucifera, and in many brown algae, e.g.Fucus vesiculosus.

D5-Avenasterol (5-Avenasterol) (�5,24Z), also called isofucosterol, 28-isofucosterol, 29-iso-fucosterol, 24Z-ethylidenecholesta-5,24(28)-dien-3b-ol, [24(28)Z]-stigmasta-5,24(28)-dien-3b-ol, or [24(240)Z]-stigmasta-5,24(280)-dien-3b-ol, C29H48O, MW 412.69, CAS # 18472-36-1.Found as a major phytosterol in oats, and in significant levels in other plant materials.

D7-Stigmasterol (7-Stigmasterol) (�7,22E) (24a=24S), also called spinasterol, (22E)-stigmasta-7,22-dien-3b-ol, C29H48O, MW 412.69, CAS # 481-18-4.

D7-Avenasterol (7-Avenasterol)(�7,24Z), also called avenasterol, (24Z)-24-ethylidenecholesta-7,24(28)-dien-3b-ol, C29H48O, MW 412.69, CAS # 23290-26-8.

In all plant tissues, phytosterols occur in five common forms (Fig. 7): as the free alcohol (FS),as fatty-acid esters (SE), as steryl glycosides (SG), and as acylated steryl glycosides (ASG). The lastthree forms (SE, SG, and ASG) are generically called ‘‘phytosterol conjugates.’’ In free phytosterols(FS), the 3b-OH group on the A-ring of the sterol nucleus is underivatized, whereas in the threeconjugates the OH is covalently boundwith another constituent. The OH group is ester-linked with afatty acid in SE and linked by a 1-O-b-glycosidic bondwith a hexose (most commonly glucose) in SG(first reported by Power and Salway in 1919 [7]). The third group of phytosterol conjugates, ASG,differ from SG by the addition of a fatty acid esterified to the 6-OH of the hexose moiety (firstreported by Lepage in 1964) [8]. Seeds of corn and rice and other grains contain a fourth type ofphytosterol conjugate, phytosteryl hydroxycinnamic-acid esters (HSE), in which the sterol 3b-OHgroup is esterified to ferulic or p-coumaric acid (Fig. 7).

2. Structural diversity and phylogenetic distribution of phytosterols

2.1. Occurrence and metabolism of cholesterol in plants

There is a widespread misconception, perhaps fostered by the nutritional information printedon packages and containers of various foods of plant origin, that plant tissues are devoid ofcholesterol. The fact is that this C27 sterol, which is predominant in animals and a contributingfactor in human cardiovascular disease, often accounts for 1–2% of the total plant sterols, andcan compose 5% or more in select plant families, species, organs, or tissues. Despite the fact thatthe edible portion of some crop plants can include cholesterol as a significant portion of the totalphytosterols, it should be noted that this is inconsequential in the human diet relative to theamount of cholesterol in meat and dairy products. Many species of the Solanaceae (Nightshade


family) include relatively high levels of cholesterol [9]. In total sterols from pericarp tissue ofmature-green tomato (Lycopersicon esculentum) fruit, cholesterol constituted �6%, and in the SEfraction it ranged from 15 to 20% [10,11]. A study of the sterol composition of seed oil from 13species of Solanum showed that in six species the combined FS plus SE fractions contained >5%cholesterol, and the level in one species, S. pseudocapsicum, ranged from 10 to 22% [12]. A recentmolecular-genetic study demonstrated that the activity of sterol methyltransferase 1 (SMT1)governs the level of cholesterol in plants [13]. SMT1 catalyzes the first step in the production ofC28 and C29 phytosterols, methylation of cycloartenol to 24-methylene cycloartenol. In matureArabidopsis plants bearing an smt1 null mutation, cholesterol was the major sterol and composed26% of the total phytosterols, compared with 6% in wild-type plants.There is ample evidence that in plants cholesterol serves as a precursor in the synthesis of steroidal

saponins and alkaloids, as well as ecdysteroids (insect molting hormones) and other pregnane- andandrostane-type steroids (see Sections 2.6, 2.7, and 2.8) [14–17]. In medicinal herbs and food plants,steroidal saponins and alkaloids are of interest because of their potential pharmacological activityand/or toxicity in animals.

2.2. Distribution and diversity of major C-24 alkyl phytosterols

The most commonly encountered phytosterols in higher plants are the 24a-methyl (24R) sterolcampesterol (Fig. 5), the 24a-ethyl (24R) sterol sitosterol (Fig. 6), and the 24a-ethyl (24S, due to

Fig. 7. Structures of phytosterol conjugates. The sites of cleavage via alkaline hydrolysis (saponification) and acid

hydrolysis are indicated with arrows.


the 22E double bond) sterol stigmasterol (Fig. 6). Each of these possesses one double bond atC5,6 in the B-ring of the sterol nucleus (�5). Typically, campesterol occurs in approximately a 2:1ratio with its 24b-methyl epimer ergost-5-en-3b-ol (22-dihydrobrassicasterol, Fig. 5) [18], whichis not readily apparent because the two isomers are not resolved by GLC or HPLC [19]. Thestereochemistry at C-24 is determined by the particular sterol methyltransferase which convertsthe initial phytosterol precursor cycloartenol (Fig. 3) to 24-methylene cycloartenol [20]. Another24b-methyl �5 sterol found predominantly in species of the Brassicaceae (also known as theCruciferae or Cruciferaceae, Mustard family) is brassicasterol (ergosta-5,22E-dien-3b-ol, Fig. 5),which generally accounts for less than 10% of the total phytosterols in crops such as cabbage,broccoli, and canola.A second major group of desmethyl sterols found in relatively few plant families includes 22-

dihydrospinasterol (7-stigmastenol, stigmast-7-en-3b-ol) and spinasterol (7-stigmasterol, stig-masta-7,22E-dien-3b-ol) as predominant constituents (Fig. 6). These are the �7 equivalents of the24a-ethyl �5 sterols sitosterol and stigmasterol, respectively, and apparently fulfill the samefunction as membrane structural components. Crops from the Cucurbitaceae (Cucumber family),including melon, squash, cucumber, and pumpkin, all contain close to 100% �7 phytosterols,whereas outside of this family the only crop plant with such a high percentage appears to bespinach, unless one includes tea as well (Camellia sinensis, a member of the Theaceae) [21].Studies early in the last decade revealed that �7 sterols are more widely distributed than pre-viously thought, and are abundant in many species from five families within the order Car-yophyllales, including the Amarathaceae, Caryophyllaceae, Chenopodiaceae, Phytolaccaceae,and Portulacaceae [21,22]. Often these species include a blend of �7 and �5 phytosterols, asexemplified by the crop plants Beta vulgaris (table beet), with a �7 to �5 ratio of about 7:3, andChenopodium quinoa (‘‘Inca wheat’’), with a �7 to �5 ratio of about 1:3 [23]. An interesting quirkof cucurbit crops is that the seeds and seedlings generally contain high levels of 24b-ethylcholesta-7,25-dienol and 24b-ethylcholesta-7,22,25-trienol, as well as small amounts of �5 phytosterols,which largely disappear as the plants grow to maturity [19,24,25].

2.3. Free and conjugated phytosterols in fruits and vegetables

The sterol lipid composition of mature-green tomato fruit is much like that of tomato leaves,and is quite unusual relative to most plant tissues and organs in that ASG accounts for more thanhalf, and ASG plus SG compose 85–90%, of the total sterols [10,26]. With tomato fruit ripening,a substantial increase in sterol synthesis was accompanied by marked changes in sterol composi-tion and conjugation [10,27]. An increase in the ratio of stigmasterol to sitosterol, the two majorphytosterols, was evident in all four sterol lipid classes but was most pronounced in FS [10,27].Although stigmasterol differs from sitosterol only by the 22E-double bond in the alkyl side chain,it has been elegantly demonstrated that these two sterols have markedly different effects on thepermeability, ordering, and fluidity of plant phospholipid vesicles [28,29]. In addition to the largeincrease in stigmasterol with ripening, there was a reapportioning of sterols among the four steryllipid classes. Both FS and SG increased at the expense of ASG, and the level of SE was 10-foldhigher in red-ripe compared with mature-green fruit [10]. A less dramatic increase in SE has beenreported to occur with senescence of both tomato leaves and potato tubers [26]. In a sterol-over-producing tobacco mutant cell line that was selected for resistance to a triazole sterol biosynthesis


inhibitor, the ‘‘excess’’ sterol was mainly metabolic intermediates that were esterified to fattyacids (SE) and sequestered in cytoplasmic lipid droplets. It was concluded that SE are involved inremoving ‘‘improper’’ sterols from the FS pool, thereby assuring ‘‘proper’’ sterol composition incell membranes [30–32]. SE from tomato fruit were also enriched in sterol intermediates and evenlate in ripening there was preferential esterification of sitosterol over stigmasterol [10].Storage of mature-green tomato fruit at 2 �C for 11–21 days (chilling) resulted in about a two-

fold increase in the level of FS. Tomatoes do not ripen at 2 �C and the increase in stigmasterolnoted with ripening of the same lot of fruit at 15 �C was greatly attenuated [11,33,34]. In micro-somal membranes from pericarp of ‘Pik-Red’ fruit stored at 2 �C, the doubling of FS was offsetby a decrease in ASG, suggesting that sterol glycosylation and esterification are inhibited at lowtemperature. This increase in the FS:ASG ratio may be a means of acclimation to low tempera-ture. The esterified fatty acids in tomato ASG are about 75–80% saturated [33,34], so a drop inthe proportion of ASG might serve to increase the fluidity of cell membranes. In accord with apossible role of ASG in thermal acclimation, a 3-day heat treatment of mature-green tomatoes at38 �C elicited a marked increase in ASG at the expense of SG and FS [35]. Four days after greentomatoes chilled for 21 days at 2 �C were transferred to 20 �C, the distribution of sterols in ASG,SG, FS, and SE had essentially returned to the pre-storage levels (63, 19, 16, and 3 mol%,respectively), but the percentage of stigmasterol had risen dramatically [34]. These fruit subse-quently showed symptoms of chilling injury, delayed and uneven ripening and extensive decay.Changes in sterol lipid content and composition in pericarp tissue during ripening of bell pepper

fruit are much less dramatic than in tomato fruit [36]. In contrast to tomato, sitosterol and cam-pesterol, in a ratio of about 3:1, are the major sterols in all four sterol lipid classes in bell pepper.Only small amounts of stigmasterol are present. FS is the most abundant of the sterol lipid classes,ranging from �55 to 75% of the total sterols in pericarp from fruit of different cultivars andseparate harvests [36,37]. The level of FS changed little with ripening and SE rose only slightly,from �5 to 7% of the total sterols. The only significant change among the sterol lipid classes,which occurred both with ripening of fruit on the plant and during a 2-week storage of mature-green fruit at 2 �C, was a 50–100% increase in SG, largely balanced by a decline in ASG [36,37].The sterol lipid distribution in microsomal membranes isolated from bell pepper pericarp tissuewas shown to change much more with ripening in the field in summer than with ripening in thegreenhouse in spring [38]. Overall, SG and ASG composed greater proportions of total sterollipids in microsomes than in whole pericarp, and a pronounced increase in microsomal SG withripening was compensated by declines of both ASG and FS. In microsomes from red-ripe field-grown fruit, SG, FS, and ASG comprised 50, 28, and 22 mol% of the total sterols, respectively.A very informative early study by Hartmann and Benveniste [39] found that a burst of

respiration and metabolic activity following slicing of potato tuber tissue is accompanied by asharp increase in de novo sterol synthesis (that utilized radiolabeled acetate). The first sterolproduct, cycloartenol, was converted to desmethyl sterols only after several hours of aging.Initially, 28-isofucosterol composed 30% of the FS fraction and pulse-chase labeling indicatedthat it was the precursor to sitosterol and stigmasterol. In retrospect, this work provided the firstindication that regulation of carbon flow into end product (24a methyl or ethyl, 4,14-desmethyl)sterols occurs at a step beyond squalene cyclization and formation of cycloartenol [3,40].As stated in Section 2.2, phytosterols of the cucurbit crops are predominantly �7, 24a-ethyl,

but also include an interesting array of unusual minor constituents. The peel and outer pericarp


tissues of zucchini squash (Cucurbita pepo) were shown to have 7-stigmasterol (spinasterol) and7-stigmastenol (22-dihydrospinasterol) as the dominant FS, each at close to 40% of the total [41].Minor sterols included 7-isoavenasterol (24E-ethylidene) at about 8%, and 7-campestenol, 8,22-stigmastadienol, sitosterol, stigmastanol, 8(9)-stigmastenol, 7,25(27)-stigmastadienol, and 7-ave-nasterol (24Z-ethylidene), all in the range of <1 to 3%. FS and SEwere present in about a 12:1 ratio;ASG and SG were not analyzed. In a postharvest study of muskmelons (Cucumis melo), the sterollipid composition was analyzed in plasma membrane (PM) isolated from hypodermal mesocarp tis-sue of mature fruit kept for either 1 night at 4 �C (pre-storage condition) or for 7 days at 4 �C plus 3days at 21 �C (post-storage condition) [42]. In PM from pre-storage melons, FS, ASG, and SGcomprised 43, 37, and 20 mol% of the total sterol lipids, respectively, whereas in PM from post-sto-rage fruit, the corresponding percentages were 57, 32, and 11. The ratio of 7-stigmasterol to 7-stig-mastenol, the two predominant sterols, was about 1.1 in FS and about 0.5 in both ASG and SG. ThePM sterol composition did not change significantly during storage in any of the sterol lipid classes.Pre-storage gamma irradiation of whole fruit at 1.0 kGy caused a transient decline in PM H+-ATPase activity, which was associated with both an increase in the proportion of FS and decreasein the 7-stigmasterol:7-stigmastenol ratio in all three sterol lipid classes. By the end of storage,however, the sterol lipid content and composition was similar in PM from control and irradiatedmelons, and H+-ATPase activity was higher in PM from the irradiated fruit.Apple (Malus domestica) fruit are the antithesis of tomato with respect to sterol conjugates, and

are surely more representative of the majority of plant tissues. At harvest, the concentrations ofFS, SG, and ASG in outer cortical tissue of ‘Golden Delicious’ apple were 145, 97, and 3 nmol/gfresh weight, respectively [43]. The mole ratio of FS:SG:ASG changed from 59:40:1 at harvest to65:33:2 after 15 weeks of storage at 0 �C plus 1 week at 20 �C, mainly due to a decline in SG from97 to 73 nmol/g fresh weight. It is interesting that a 4-day heat treatment at 38 �C just after har-vest also induced a specific reduction in SG to 77 nmol/g fresh weight. The level of ASG wasincreased after a longer duration of cold storage (6 months at 0 �C), but the concentrations of FSand SG remained about the same [44]. The sterol composition of both FS and SG included 90–95% sitosterol and changed little with storage. Minor sterols in the FS fraction were identified ascampesterol, stigmasterol, 5,7-stigmastadienol, and 5,25(27)-stigmastadienol [43].Carrot (Daucus carota) storage root tissue appears to respond to wounding in much the same

way that potato tuber tissue does, i.e. there is an induction of de novo phytosterol synthesisassociated with cell membrane proliferation and repair [45]. However, tissues from the two sto-rage organs do differ markedly with respect to enzymatic hydrolysis of glycerolipids directly afterwounding, which is extensive in potato but minimal in carrot [46]. In a pair of studies, freshlyshredded carrots were stored at 10 �C and 95% relative humidity for up to 10 days, and sampleswere taken periodically for analysis of membrane sterol lipid and glycerolipid contents [45,47].After 10 days of storage, total sterol lipids (FS+ASG+SG) in the shredded tissue had increasedby 24–28%. In the first study, the mole ratio of FS:ASG:SG in the carrot tissues changed from76:17:7 immediately after shredding to 64:30:6 after 10 days, whereas in the second study thechange was from 56:29:15 to 60:33:7. The tissue concentrations of ASG and FS increased in bothstorage experiments, whereas SG remained the same or declined. In the FS fraction, sitosterolcomposed 56–68%, stigmasterol 16–28%, and campesterol 10–12%. The only significant changein FS composition with storage was an increase in the stigmasterol:sitosterol ratio from 0.30 to0.45 during the last 5 days at 10 �C.


2.4. Unique phytosterols and phytosterol conjugates in cereals

Several unique phytosterols and phytosterol conjugates have been reported in cereal grains.Seitz [48] reported trans-hydroxycinnamate esters of phytosterols (HSE, including steryl ferulateand p-coumarate esters) of phytosterols in corn, wheat, rye, rice, and triticale. Norton [49,50]extended these studies and separated several additional molecular species of hydroxycinnamateesters from rice bran and corn bran. We reported that the levels of hydroxycinnamate esters incorn fiber were higher than in corn bran or any other grain [51]. In a recent paper, we comparedthe levels of SE, FS, and HSE in 66 accessions of Zea, teoscinte and Job’s tears, and identifiedseveral corn accessions with very high levels of HSE and total phytosterols [52]. Both Seitz andNorton noted that sitostanol was the predominant phytosterol in corn HSE [48,50], whereascycloartanol and 24-methylene cycloartenol were the predominant phytosterols in rice bran HSE(called ‘‘oryzanol’’). Piironen and colleagues [53] recently compared the composition of totalsterols (free+bound) in rye, oats, barley, wheat, corn and other grains. They found that all grainscontained significant levels of phytostanols (sitostanol and campestanol) in the total phytosterolfractions. Recent studies from our lab indicate that most of the phytostanols in corn are esterifiedin either SE or HSE, and all of the HSE is localized in the aleurone cells which form a single layerin corn, and fractionates into the corn fiber fraction during wet milling [54]. Since commercialcorn oil is obtained by extracting corn germ, the levels of HSE and phytostanols in corn germ oilare very low [55,56].

2.5. Function of 24-alkyl phytosterols and their conjugates

In plant as in animal cells, the plasma membrane is greatly enriched in sterols relative to othercell membranes [57,58]. The profound effects of sterols on the physical properties of membranesare well documented [59]. Through interaction with phospholipids in a one to two stoichiometry,sterols condense the bilayer, reduce bulk fluidity and permeability, and broaden or eliminatephospholipid phase transitions [60,61]. Sterol–phospholipid interactions influence membranefunctions such as simple diffusion, carrier-mediated diffusion, and active transport, and alsomodulate the activities of membrane-bound enzymes or receptors [62]. Although a wide variationin sterol structure can be accommodated to fulfill the bulk membrane structural requirement [59],there appear to be other, more subtle functions or specific situations, such as salt stress, for whichsterol structure becomes more critical [63,64]. Evidence from sterol biosynthesis inhibitor studiesindicate that phytosterols also play an essential role in plant cell division [65–67].Compared with work on membrane phospholipids, the role of sterol lipids in pre- and post-

harvest plant physiology has received little attention [3,68,69]. A sharp increase in the ster-ol:phospholipid ratio in microsomal membranes during plant senescence is associated with loss ofmembrane function [70,71]. Changes in sterol composition likely to affect membrane function[28,29,57] can occur with greening, shading, maturation, aging, or ripening of plant tissues [10,72–74] . There is evidence that free sterols are tightly bound to the plasma membrane H+-ATPaseand may be essential for activity of this critical enzyme [75]. Further work has shown that H+-ATPase activity is dependent upon the kind and amount of sterol present in a reconstituted sys-tem [76,77]. Finally, Zelazny and colleagues [78] showed that free sterols in the plasma membraneof the marine alga Dunaliella are absolutely required for sensing osmotic changes.


Sterol conjugation, the conversion of free sterols (FS) to steryl esters (SE), steryl glycosides(SG), or acylated steryl glycosides (ASG), is another potentially important aspect of membranelipid metabolism. Like FS, SG and ASG are membrane structural components, whereas SEappear to be largely excluded from membranes [31,79], possibly because of their relatively lowsolubility in a phospholipid bilayer [80]. Metabolic studies have shown that interconversion ofsterols and sterol conjugates is quite rapid, suggesting a regulatory function [81]. Based on reportsthat sterol interconversions are controlled by phytohormone levels and environmental factorssuch as light, temperature, and water stress, it has been postulated that they are involved in theregulation of membrane properties in response to changing conditions [62]. Kesselmeier and col-leagues [82] reported an increase in the levels of SG and ASG during the enzymatic preparationof protoplasts from oat leaves. In the course of our work, we have found that the extent of sterolglycosylation and esterification can be dramatically altered in response to ozone stress in snap-bean leaves [83], growth conditions in bell pepper fruit [38], freezing stress in potato leaf plasmamembrane [84], chilling stress in tomato fruit [34], and challenge by fungal elicitors, cellulase,xylanase, or copper ions in tobacco cells [85]. In particular, it appears that different types of stresscan promote conversion of FS to ASG [82,83,85]. Recently, Peng and colleagues [86] reportedthat sitosterol-b-glucoside serves as a primer for cellulose synthase in plants. Since it is thoughtthat most of the ASG and SG is localized in the plasma membrane [84,85], the involvement of SGin a biosynthetic process that occurs adjacent to the plasma membrane is a reasonable hypothesis.

2.6. Steroidal saponins

Steroidal saponins consist of a furostanol- or spirostanol-based aglycone (Fig. 8) and an oli-gosaccharide of typically 2–5 hexose or pentose moieties attached to the 3b-OH of the steroidnucleus. These saponins are abundant in a number of monocot species, including the crops onion,garlic, and leek (Allium spp.), Asparagus, oats (Avena sativa), and yam (Dioscorea spp.), and havealso been described in several solanaceous crops, including eggplant (Solanum melongena), bellpepper (Capsicum annuum), and tomato, and the leguminous herb fenugreek (Trigonella foenum-graecum) [16, 87–90]. Glucosylation of yamogenin (25S-spirost-5-en-3b-ol) at the 3b-OH by asteroid-specific UDP-glucose-dependent glucosyl transferase, the first step in generation of a ster-oidal saponin from the sapogenin aglycone, was demonstrated in a preparation from Asparagusplumosus [89]. Also, the furostanol saponins are usually present as 26-O-glucosides that are cleavedby specific 26-O-b-glucosidases. In oats, one such enzyme converts avenacosides A and B toantifungal 26-desglucoavenacosides [87], whereas another 26-O-b-glucosidase cloned from Costusspeciosus (wild ginger) transforms furostanol saponins lacking the F-ring to the correspondingspirostanol saponins (Fig. 8) [91].Variations in sapogenin spirostan and furostan structures (e.g. addition of hydroxy groups at

C-2 and/or C-22, and saturation versus unsaturation at C5,6), and the myriad possible arrange-ments of oligosaccharides glycosylated to the 3-b-OH, allow for many different steroidal saponins.Dozens have been isolated from the monocot and dicot crops cited above, including at least 17 fromAsparagus spp., 10 from leek, and eight from fenugreek [87,88,90]. Among these, a number are ofpharmacological interest, such as porrigenin C, which showed antiproliferative activity on fourtumor cell lines [90], and a spirostanol glycoside from fruit of Asparagus officinalis with markedspermatocidal activity [88]. Yams, wild alliums, and fenugreek have drawn much attention as rich


sources of diosgenin and yamogenin (Fig. 8), which are used in production of various steroids.Finally, preparations of steroidal saponins from Dioscorea, fenugreek, and particularly the Indianpuncture plant, Tribulus terrestris, are currently widely marketed on the Internet as herbal sub-stitutes for anabolic steroids and ViagraTM because of their reported stimulation of testosteroneproduction.

2.7. Steroidal glycoalkaloids

Steroidal glycoalkaloids appear to be ubiquitous in members of the Solanaceae and among thesolanaceous crops they are most abundant and diverse in wild and cultivated potato (Solanumspp.) [15,87,92]. The steroidal alkaloid aglycones have solanidane- or spirosolane-based struc-

Fig. 8. Steroidal sapogenins (aglycones) and saponins.


tures, the latter being closely related to the spirostanol sapogenins [92]. The common solanidaneaglycones include solanidine (solanid-5-en-3b-ol) and demissidine (5a-solanidan-3b-ol), andcommon spirosolane aglycones include solasodine (22R,25R-spirosol-5-en-3b-ol), soladulcidine(22R,25R,5a-spirosolan-3b-ol), tomatidenol (22S,25S-spirosol-5-en-3b-ol), and tomatidine(22S,25S,5a-spirosolan-3b-ol) (Fig. 9). Two trisaccharides of solanidine, a-solanine (3b-O-galac-tose-rhamnose1glucose2) and a-chaconine (3b-O-glucose-rhamnose1rhamnose2), are the mainsteroidal glycoalkaloids in S. tuberosum. Two analogous trisaccharides of tomatidenol, a- andb-solamarine, respectively, as well as two tetrasaccharides of demissidine, have been introducedinto cultivated potato by crossbreeding with wild species [87,92]. In tomato, tomatidine is theprincipal aglycone and its tetrasaccharide a-tomatine (3b-O-galactose-glucose-xylose1galactose2)is the major glycoalkaloid, whereas the major constituents in eggplant are the aglycone solasodineand its trisaccharide solasonine (3b-O-galactose-rhamnose1glucose2) [93]. Accumulation of sola-nine and chaconine in potato tubers is closely associated with light-induced greening (chlorophyllsynthesis), although metabolically these appear to be independent events [94,95]. Similarly, intomato fruit a-tomatine accumulates in immature green fruit but is essentially absent in fully ripefruit. Radiolabeling experiments showed that young tomato fruit synthesized a-tomatine fromcholesterol and that the decline in a-tomatine concentration with ripening could be attributed toloss of biosynthetic capacity combined with an increased rate of degradation [96].The steroidal glycoalkaloids are toxic to humans and other mammals. Fortuitously, they are

poorly absorbed by the gastrointestinal tract, where they are partially hydrolyzed to the less toxicaglycones, which in turn are rapidly excreted [97]. At moderately high concentrations (�20 mg/100 g fresh weight), these alkaloids have a bitter taste and create a burning sensation in the mouthand throat. They are intense irritants of the gastrointestinal tract due to disruption of cell mem-branes, and also act as cholinesterase inhibitors, which can severely depress the central nervoussystem [94,95,97,98]. A toxicological study in which four glycoalkaloids or their respective agly-cones were fed to mice indicated that a-chaconine is more damaging to the liver than a-solanine,solasonine, or a-tomatine, and although solanidine and solasodine (but not tomatidine) caused

Fig. 9. Steroidal aglycones commonly found in glycoalkaloids of the Solanaceae.


significant liver enlargement, this was reversible upon removal of the alkaloids from the diet [93].A recent extension of this work determined that among the aglycones, tomatidine, with a satu-rated steroid nucleus, is essentially non-toxic relative to the 5-unsaturated solanidine and solaso-dine [99]. On the other side of the ledger, there have been a number of reports indicating thatsome steroidal alkaloids have potential pharmacological or therapeutic applications [98]. Forexample, solasodine has hypocholesterolaemic and antiatherosclerotic effects [100], whereas itsglycosides (solasonine and solamargine) show selective toxicity against cancer cells [101].

2.8. Phytoecdysteroids and brassinosteroids

Phytoecdysteroids, analogues of insect-molting hormones (ecdysteroids) that play an essentialrole in insect development and maturation, are found in higher plant species representing over100 families [23,102]. More than 150 different phytoecdysteroid structures have been reported[103], the most commonly encountered being 20-hydroxyecdysone (Fig. 10), which is identical to theprincipal molting hormone isolated from insects [23]. Although definitive proof is still lacking, it iswidely accepted that production of phytoecdysteroids in plants deters predation by non-adaptedinsects (as well as other invertebrates such as nematodes), either by functioning as antifeedants or viadisruption of development [102].The occurrence, biosynthesis, and distribution of these plant steroids have been studied most

extensively in species of the Chenopodiaceae (Goosefoot family). Accumulation of phytoecdys-teroids produced in young, developing tissues generally occurs in apical leaves and flowers andultimately in seeds [104,105]. Appreciable levels were detected in seeds of about 35% of the spe-cies tested [102]. Among chenopod crop species, leaves and seeds of spinach (Spinacia olereacea)are particularly rich in phytoecdysteroids and levels are also high in Chenopodium quinoa (theInca ‘‘mother grain’’ currently being developed as a dryland crop in the western USA), whereas intable beet (Beta vulgaris) these steroids are scarcely detectable [23,102].The biosynthetic origin of phytoecdysteroids is an interesting and as yet incomplete story. A

number of studies have shown incorporation of radiolabeled cholesterol into these plant steroids,leading to the conclusion that cholesterol serves as the sterol precursor [9,15]. However, in spi-nach, which produces exclusively �7 rather than �5 sterols (see Section 2.2), it has been shown

Fig. 10. Phytoecdysteroid—a plant steroid that serves as an insect molting hormone. 20-Hydroxyecdysone is synthe-sized from lathosterol in spinach (Spinacia oleracea) and is identical with the naturally-occurring hormone in insects.


that lathosterol (cholest-7-en-3b-ol) is the direct precursor of 20-hydroxyecdysone [106]. Whetherthere are separate biosynthetic routes to phytoecdysteroids utilizing cholesterol and lathosterol,and whether cholesterol is converted to lathosterol in some plants by consecutive hydrogenationand dehydrogenation steps (Fig. 4), are presently open questions [23]. Aside from this issue, it isalso known that many plants produce ecdysteroids that are alkylated at C-24 and thus are likelyderived from C-24 methyl and ethyl phytosterols [107].In the last decade, brassinosteroids have come to be recognized as an important new class of

potent steroid hormones in plants, operative in the nanomolar range or lower as are their steroidcounterparts in animal cells [108,109]. Over 40 of these plant steroids have been fully character-ized, the most active compound being brassinolide (Fig. 11), which was also the first to be iso-lated [110]. Mutants that are deficient in the synthesis of or responsiveness to brassinosteroids aretypically characterized by a dwarf phenotype when grown in the light and deetiolation whengrown in the dark, and are also often impaired in reproduction [108–110].The complete, dual biosynthetic pathways of brassinolide in Catharanthus roseus and Arabi-

dopsis have recently been elucidated through detailed metabolic studies in which deuterium-labeled intermediates were supplied to cultured cells of C. roseus and various Arabidopsis mutants[109–111]. Campesterol is the mainstream desmethyl phytosterol from which brassinolide and itsimmediate precursor castasterone (Fig. 11), another active brassinosteroid, are derived. It wasrecently determined that the brassinolide-deficient Arabidopsis mutant dim (also called dwf1) lacksthe ability to convert 24-methylenecholesterol to campesterol via a 24-methyldesmosterol inter-mediate [109]. An interesting observation from this study, unrelated to brassinolide synthesis, wasthat dim mutant plants accumulate very high levels of isofucosterol, indicating that the DIM geneproduct also performs the reduction of isofucosterol to sitosterol via the stigmasta-5,24(25)-die-nol intermediate. The initial steps in the dual pathways from campesterol to castasterone andbrassinolide involve the conversion of campesterol to campestanol. This is actually a three-stepprocess that includes 24-methylcholest-4-en-3-one and 24-methyl-5a-cholestan-3-one as inter-mediates. A recent study of the brassinolide-deficient Arabidopsis mutant det2 showed that theDET2 gene product saturates the 4-double bond in the A-ring of 24-methylcholest-4-en-3-one to

Fig. 11. Brassinosteroids: a recently discovered class of plant hormones derived from campesterol and other phytosterols.


yield 24-methyl-5a-cholestan-3-one [111]. In the subsequent metabolism of campestanol to cas-tasterone, there are two alternate routes dubbed the ‘‘early’’ and ‘‘late’’ C6-oxidation pathways,which refers to whether the 6-keto group present on the B-ring of castasterone is introducedduring the first step (campestanol to 6-oxocampestanol) or during the last step (6-deoxocastaste-rone to castasterone) [110]. It is interesting that the products of two genes in Arabidopsis, DWF4and CPD, are cytochrome P450-like enzymes that perform the sequential hydroxylations at C22and C23 of the alkyl side chain in both the early and late C6-oxidation pathways of brassinolidebiosynthesis [109,110].

3. Methods for the quantitative analysis of phytosterols and phytostanols

3.1. Extraction and fractionation of phytosterols

Most common methods for the extraction of lipids also extract phytosterols. Nonpolar solventssuch as hexane (commonly used to extract most types of vegetable oils), quantitatively extractfree phytosterols (FS) and phytosteryl fatty-acid esters (SE) [1,3]. The extraction of FS, SE, andferulate phytosteryl esters from corn fiber was compared with four different solvents (hexane,methylene chloride, ethanol, and isopropanol) and each solvent extracted >95% of these threesterol lipid classes [51]. Steryl glycosides (SG) and fatty-acylated steryl glycosides (ASG) are onlypartially extracted with hexane, and increasing the polarity of the solvent gave a higher percen-tage of extraction [1]. We routinely use the Bligh and Dyer chloroform–methanol extractionmethod to extract all sterol lipid classes [112]. Even after chloroform-methanol extraction, addi-tional phytosterols are sometimes liberated by subsequent acid or alkaline hydrolysis, suggestingthat there may be pools of ‘‘bound’’ or ‘‘evasive’’ phytosterols in some plant tissues (personalcommunication, V. Piironen). Additional research is necessary to provide a better understandingof the optimal extraction methods that are required to assure complete phytosterol extraction.Once a lipid extract has been prepared it is often necessary to separate (fractionate) one or more

of the sterol lipid classes. Traditionally, ‘‘open column’’ LC (liquid chromatography), with eithersilicic acid or Floricil as the solid phase column packing, was used to fractionate the various lipidclasses. In the last decade, open column LC has generally been replaced by similar methods usingpre-packed SPE (solid phase extraction) cartridge columns. We recently reported a silica SPEmethod to purify hydroxycinnamate steryl esters (HSE) in corn fiber oil [113]. In a later sectionwe will present several SPE methods for the fractionation of free phytosterols before GC analysis.Preparative thin-layer chromatography can be used to separate and fractionate sterol lipid clas-ses. After spotting the sample(s) on a TLC plate, the plate is developed with an appropriate sol-vent mixture, and each ‘‘spot’’ containing a specific sterol lipid class is scraped into a tube andeluted with solvent. Finally, silver ion (‘‘argentation’’) chromatography (either silver-impregnatedTLC or HPLC) can be employed as a fractionation technique that separates phytosterols basedon their total number of carbon–carbon double bonds [1].Before the development of micro-scale GLC and HPLC analytical methods, precipitation of

cholesterol with digitonin (a type of steroidal saponin) was a technique commonly used to removeother lipids and obtain a cholesterol-enriched fraction. Digitonin precipitation also has been usedin studies of both fungal sterols [114] and phytosterols [115] to isolate the FS fraction (binding of


digitonin to a sterol, and consequent precipitation of the complex, requires a free 3b-OH group).However, there is evidence that certain types of phytosterols are not quantitatively precipitatedby digitonin, and this technique is now infrequently used to isolate free phytosterols.

3.2. Methods for the separation of intact phytosterol classes

Thin-layer chromatography was traditionally used for qualitative separation of phytosterollipid classes. Grunwald and Huang [116] compared five different TLC methods for separation ofthe four common sterol lipid classes in plant tissues (FS, SE, SG, and ASG). Numerous otherTLC methods for separation of phytosterol lipid classes have been reported [1,3].Numerous high performance liquid chromatography (HPLC) methods have been developed to

both qualitatively separate and quantitatively analyze phytosterol lipid classes (Table 1). In gen-eral, polar or ‘‘normal phase’’ HPLC columns (e.g. silica, DIOL, Amino, CN) are used to separatephytosterol lipid classes (Table 1) [117–123] and ‘‘reversed phase’’ HPLC columns (e.g.C18=ODS, C8, or phenyl) are used to separate molecular species (individual compounds) thatcomprise a lipid class (see next section and Table 2). Although methods have been reported forthe simultaneous analysis of both nonpolar (FS, SE, and sometimes HSE) and polar (SG andASG) sterol lipid classes [119,121], we find that the most accurate way to quantify these lipids isto analyze the polar and nonpolar classes separately (Figs. 12 and 13) [85,120]. In our metho-dology, the filtered total lipid extract is injected in two different HPLC systems, both equippedwith a Diol column but programmed for different solvent gradients. The three nonpolar sterollipid classes are quantitatively analyzed with a hexane-based gradient that starts at 0% iso-propanol and increases to 0.25% during 40 min (Fig. 12), resulting in retention times of 2, 21, and28 min for SE, FS, and HSE [51]. The two polar sterol lipid classes are quantitatively analyzedwith a gradient that starts at 90/10, hexane/isopropanol, and increases to 45/50/5, hexane/iso-propanol/water, resulting in retention times of 6 and 11 min for ASG and SG, respectively, withthe other glycolipids and phospholipids eluting over the range of 15–50 min (Fig. 13).

Table 1Methods for the TLC and HPLC analysis of intact phytosterol classes

Sample Chromatography Gradient SE HSE FS ASG SG Date Reference

Standards and various plants TLC-Silica Y Y Y Y 1989 [116]

Fruit and vegetables TLC-Silica Y Y 2001 [117]Peanut and corn oils TLC-Silica Y 1984 [118]Peanut and corn oils Low pressure LC-Silica Steps Y Y 1984 [118]Spinach leaves HPLC-Silica Y Y Y Y Y 1990 [119]

Tobacco cells HPLC-Cyano Y Y Y Y 1994 [120]Wheat flour HPLC-Silica Y Y Y Y Y 1993 [121]Corn fiber HPLC-DIOL Y Y Y Y 1996 [51]

Rice bran oil HPLC-Prep Nova-PakHR Silica

Y 1999 [122]

Zooplankton lipids HPLC-Alumina Y Y 1999 [123]

Soy lecithin HPLC-DIOL Y Y Y Fig. 9 Unpublished

SE, fatty acid ester; HSE, hydroxycinnamic-acid esters; FS, free alcohol; ASG, acylated steryl glycoside; SG, sterylglycoside.


3.3. Methods for the separation of molecular species of phytosterol conjugates

When the five phytosterol lipid classes (FS, SE, HSE, SG, and ASG) have been separated byTLC or HPLC, it is sometimes useful to study the individual compounds (molecular species)within each class. Billheimer and colleagues [124] reported the first reversed phase HPLC methodto separate molecular species of SE (stigmasteryl oleate and campesteryl palmitate are twoexamples of SE molecular species). Kesselmeier and colleagues [125] reported a method to sepa-rate molecular species of FS and SG, and also stated that the method could be used to analyzeSG molecular species obtained by partial hydrolysis of ASG (selective cleavage of the fatty acid

Table 2Methods for the HPLC analysis of molecular species of free phytosterols and intact phytosterol conjugates

Sample Column SE FS HSE ASG SG Date Reference

Synthetic standards Zorbax ODS Y 1983 [124]

Oat leaves and seeds RP-hexyl Y Y 1985 [125]Fruits and vegetables Luna C18 HPLC Y Y 2001 [117]Grains Zorbax C18 Y 1989 [48]Corn and rice Deltabond C18 Y 1995 [50]

Rice bran oil Microsorb-MV C18 Y 1999 [122]

SE, fatty acid ester; HSE, hydroxycinnamic-acid esters; FS, free alcohol; ASG, acylated steryl glycoside; SG, sterylglycoside.

Fig. 12. HPLC Chromatogram of nonpolar lipids including SE, FS, and HSE, using a DIOL column and detection viaan evaporative light scattering detector [51].


from the 6-OH of the hexose moiety). A recent report described an HPLC method to separateintact molecular species of ASG and SG, as well as the three other common plant glycolipidclasses [117]. Several methods have been reported for separating molecular species of HSE fromcorn fiber oil, rice bran oil, and lipids of other grains (Table 2) [124,125]. The most abundantmolecular species of HSE in corn fiber oil and rice bran oil are sitostanyl ferulate and cycloartanylferulate, respectively [50].

3.4. Methods for hydrolysis of conjugates and separation of free phytosterols

Total phytosterols (including free, esterified and glycosylated) can be quantified by hydrolysisand subsequent GC analysis of the combined FS. This method of phytosterol analysis has beenthe most widely used. SE, HSE, and the fatty acid-hexose ester linkage in ASG can be hydrolyzedvia saponification (alkaline hydrolysis with 1–2 N KOH or NaOH), whereas the glycosydic lin-kages in SG and ASG require acid hydrolysis (4–6 N HCl). The points of hydrolytic cleavage ofthe four phytosterol conjugates are indicated in Fig. 7. Since vegetable oil samples usually containprimarily SE and little or no SG or ASG, alkaline hydrolysis alone is sufficient to cleave all of theconjugated phytosterols [55]. Toivo and colleagues [126,127] developed an elaborate, routinemethod for hydrolysis of phytosterol conjugates that includes both acidic and alkaline steps.However, these authors noted that additional studies would be necessary to optimize the condi-tions for acid hydrolysis of phytosterol conjugates. One artifact observed with this method is thatif plant tissues contained �5-avenasterol, acid hydrolysis caused its isomerization to fucosteroland several 5,23- and 5,24(25)-stigmastadienols [128]. Similarly, we found that during acidhydrolysis of SG, �5-avenasterol (isofucosterol) is ‘‘lost’’ and lathosterol (cholest-7-en-3b-ol),included as an internal standard, was isomerized to cholesterol (B.D. Whitaker, unpublishedobservation). Kamal-Eldin et al. [128] suggested that replacing acid plus alkaline hydrolysis pro-tocols with enzymatic hydrolysis would solve this problem. Kesselmeier and colleagues [125]reported an enzymatic method (using a commercial b-glucosidase) to hydrolyze SG, but this

Fig. 13. HPLC Chromatogram of polar lipids including ASG and SG, using a DIOL column and detection via anevaporative light-scattering detector (R. Moreau, unpublished results).


method has been used by only a few researchers. We used a modified version of the enzymaticmethod [125] to analyze FS derived from SG (and ASG by including mild alkaline methanolysis,which also enabled us to identify ASG fatty acids by GC analysis of their methyl esters) [35,83–85]. The enzyme ‘‘cholesterol ester hydrolase’’ is used to hydrolyze cholesteryl fatty-acid estersduring the routine enzymatic assay of cholesterol (see Section 3.6), and could perhaps also beused to hydrolyze phytosteryl fatty-acid esters. Although cholesterol esterase has been used tohydrolyze carotenoid esters [129], we are unaware of studies employing it to hydrolyze phytos-teryl fatty-acid esters. Clearly, more research is required to perfect hydrolysis methods for phy-tosterols, and enzymatic methods may prove to be superior to acidic and/or alkaline hydrolyses.After hydrolysis of conjugates, the quantitative GC analysis of FS can be achieved using one of

the numerous published methods (Table 3) [130–136]. Most GC methods for phytosterol analysisinclude derivatization to form either trimethylsilyl (TMS) ethers (using BSTFA or a related sily-lating reagent) or phytosteryl acetates (via acetylation with pyridine and acetic anhydride), butsome methods show good separation and quantification of underivatized phytosterols (Fig. 14).Although quantitative GC analysis of underivatized phytosterols appears to be accurate andreliable, some experts prefer derivatizing phytosterols to prevent dehydration and decomposition,which may result in peak tailing and poor resolution [1]. Several reversed phase HPLC proce-dures for the separation of FS have also been reported [1,3,125] but the separation and sensitivityare generally lower than with GC.

Table 3

Methods for the hydrolysis and GC analysis of free phytosterols (unless otherwise noted, all columns were capillarycolumns)

Sample Alkalinehydrolysis

Acidhydrolysis

Enzymatichydrolysis

Derivatization SPEa Column Date Reference

Oat leaf and seeds Y Y Y None OV-1 1985 [125]

Cucumber Acetylation OV-17 1987 [19]Tomato fruit None SP2100,

packed column1988 [10]

Bell pepper Y None SPB-1 1989 [130]Yeast Y Silylation SPB-1 1989 [131]Tobacco cells Y Y None SPB-1 1994 [85]Vegetable oils Y Silylation OV-1 1996 [132]

Oat seeds Y Y Silylation DB-5ms 1998 [128]Fruit juices None DB-5ms 1998 [133]Vegetable oils Y Silylation Y NB-17 1998 [134]

Vegetable oils Y Silylation Y DB 17 HT 1999 [135]Oat seeds Y Y Silylation DB-5ms 1999 [136]Vegetable oils Y None Y SAC-5 2000 [55]

Grains Y Y Silylation Y NB-17 2000 [126]Corn fiber oil Y None SAC-5 2000 [54]Grains Y Y Silylation Y RTX-5w/

INTEGRA2002 [53]

a SPE, solid phase extraction.


3.5. Mass spectrometry and NMR of phytosterols

Mass spectrometry (MS) and nuclear magnetic resonance spectroscopy (NMR) are valuabletools for phytosterol identification. Goad’s review [1] contains an excellent comprehensive dis-cussion of these topics and it also contains a listing of MS electron impact ion fragments andNMR resonances for all of the common phytosterols. We will present a broad overview of thesetopics but would urge the reader to consult previous reviews [1] for more details.Electron impact (EI) GC–MS has been extensively employed to identify free phytosterols. Sev-

eral common free phytosterols are included in two ‘‘on-line’’ databases: http://webbook.nist.gov/chemistry and http://lipid.bio.m.u-tokyo.ac.jp. One complicating factor when employing GC–MSfor identification of phytosterols is the fact that published spectra include data for free phytos-terols (Fig. 15) and for their TMS-ether and acetate-ester derivatives. The several common pointsof EI–MS fragmentation (Fig. 15) can be used to predict fragments in phytosterols with otherstructures. Rahier and Benveniste [137] described the mass spectra of six common phytosterolswhich were each analyzed by GC–MS as the free form (M+), as acetate esters (M++42), or asTMS ethers (M++72). Because the fragmentation pattern of phytosterols is different when theyare analyzed in each of these three forms, accurate structural identification is possible only if thespectrum of the unknown compound is matched with a library spectrum of a known compoundin the same form (i.e. free, acetate, or TMS). Since many of the common phytosterols occur asepimers (e.g. brassicasterol and epibrassicasterol) or isomers (e.g. �5-avenasterol and fucosterol),GC separation and/or identification of these structurally similar compounds can be challenging.In a recent study reporting the acid-induced isomerization of �5-avenasterol and fucosterol [128],small differences in the GC-MS fragmentation of TMS derivatives were used to document thisisomerization. Another study [138], employing GC–MS of phytosteryl acetates, showed that�5-avenasterol (24Z) always has a longer GC retention time than fucosterol (24E), and that thesetwo isomers can be distinguished by the mass spectra of their acetates. M-60=m/e 394 is muchmore abundant in the fucosterol EI spectrum, whereas m/e 296 is much more abundant in the

Fig. 14. GC–MS chromatogram of sterols and stanols in saponified corn fiber oil (R. Moreau, unpublished results).


�5-avenasterol spectrum. Unique EI–MS fragments of 24E- compared with 24Z-ethylidenephytosteryl acetates were also reported elsewhere [139].Another study employed HPLC–MS, specifically atmospheric pressure chemical ionozation mass

spectroscopy (APCI–MS), to provide structural information about SG and ASG in red bell pepperfruit [117]. Other emerging forms of ‘‘soft’’ ionization mass spectrometry (e.g. electrospray andMALDI-TOF) could also potentially be used to analyze phytosterols and their intact conjugates.

13C nuclear magnetic resonance (NMR) has been reported to be a valuable tool for the chemicalidentification of phytosterols since each of the 28 or 29 carbons in the molecule can potentially beexamined individually [140]. 1H and 13CNMRhave proven to be essential in the structural elucidationand quantification of 24R- and 24S-epimers of C-24 methyl and ethyl phytosterols [1,19].

3.6. Enzymatic assays of phytosterols

Enzymatic methods [141] and specific test kits have been developed and marketed to measurecholesterol in blood and other samples. Most of these kits include cholesterol ester hydrolase tohydrolyze cholesteryl fatty-acid esters and cholesterol oxidase (from animal or microbial sources)

Fig. 15. Electron impact mass spectrum of sitosterol (MW 414) and its major fragment ions (spectrum was kindlyprovided by A. Nunez).


to oxidize cholesterol, and for each molecule of cholesterol that is oxidized, one molecule of H2O2

is produced (Fig. 16) [142]. The amount of H2O2 produced can then be measured by one of sev-eral spectrophotometric or fluorometric assays and used to calculate the amount of cholesterol inthe sample. Although the manufacturers of some of these kits indicate that the kits can also beused to measure phytosterols, we are not aware that this claim has been validated. Smith andBrooks [143] compared the Km and Vmax values for the oxidation of several phytosterols bycholesterol oxidase from Nocardia erythropolis (Fig. 16), and found that the values varied con-siderably. Obviously, with the current interest in phytosterols, there is a potential to developenzymatic methods for the rapid quantification of phytosterols in foods. When one considers thehigh cost of GC and HPLC instruments that are currently required to analyze phytosterols, thedevelopment of convenient enzymatic assays using microtiter plates is very attractive.

4. Health-promoting effects of phytosterols, phytostanols, and their esters

4.1. Historical perspectives, changing dogma, and critical questions

Phytosterols are natural components of human diets. In the West, we consume an average of�250 mg per day of phytosterols, largely derived from vegetable oils, cereals, fruits and vege-tables [144,145]. This is roughly equivalent to the amount of cholesterol (�300 mg/day) con-sumed. For vegetarians, dietary phytosterols have been estimated at almost twice this level [146].Phytostanols are much less abundant in nature than phytosterols, and consequently we typicallyconsume much lower amounts (�25 mg/day) in our diets [145,146]. Common dietary sources ofphytostanols are corn, wheat, rye, and rice. For many years, the existence and dietary effects ofthese minor sterols were largely ignored and poorly understood. Sterol chemists and biochemistsfocused their efforts on cholesterol because elevated serum cholesterol levels were shown to be aprominent risk factor for cardiovascular disease (CVD). Recent strategies for lowering serum

Fig. 16. Cholesterol oxidase reaction with cholesterol and stigmasterol. Note that oxidation of both sterols shifts the�5 double bond to �4. Enzyme kinetic data were reported for cholesterol oxidase from Nocardia erythropolis [143].


cholesterol (and risk of CVD) utilize dietary restrictions to limit cholesterol intake and/or requirethe use of drugs such as the popular ‘‘statins’’ which inhibit cholesterol biosynthesis in humans.The prospect of lowering cholesterol levels by consuming ‘‘functional’’ foods fortified with nat-ural phytonutrients would seem more attractive to many than use of drugs or dietary restrictions.Since the mid 1990s, there has been considerable interest and commercial marketing of phy-

tosterol products for this purpose. Contrary to what newcomers to the field might think, the useof phytosterols to lower serum cholesterol levels is not a new idea. Sitosterol, for instance, wasused in the 1950s [147] as a supplement and as a drug (Cytellin, marketed by Eli Lilly) to lowerserum cholesterol levels in hypercholesterolemic individuals (see [3] for an excellent review ofearly research and clinical studies on phytosterols). Due to poor solubility and bioavailability ofthe free phytosterols, the serum cholesterol-lowering effects were not always consistent, and veryhigh doses (up to 25–50 g per day) were sometimes required for efficacy. This problem of solubi-lity and bioavailability led to many confounding results in early clinical studies, and when the morepredictable and effective ‘‘statin’’ drugs became available, the use of these phytosterol productsdiminished rapidly. Thus, for a time, phytosterols were considered to be of little practical interest forserum cholesterol management. In recent years, with the growing interest in functional foods, the useof phytosterols for reducing serum cholesterol levels has regained considerable momentum. This canbe attributed, in large part, to the work of scientists at Raisio Group in Finland, who esterified phy-tostanols with fatty acids (stanyl esters) in order to improve their solubility and this allowed the firstpractical, commercial scale production of phytosterol-containing foods such as margarines [148].Procter & Gamble demonstrated the enhanced solubility of phytosterol fatty acid esters in fattyfoods such as cooking and salad oils in the early 1970s [149] but the Raisio group was the first touse food-grade esterification methods to successfully prepare and commercialize phytostanylester-containing margarines that were demonstrated in clinical studies (see Section 4.2) to con-sistently lower LDL-cholesterol levels at low (2–3 g/day) dosage levels.Because of the success with esterified forms of stanols, many researchers lost interest in free

phytosterols and phytostanols and it appeared that stanyl esters were superior to all other sterolforms for cholesterol reduction. However, subsequent research has shown that fatty acid esters ofsterols (steryl esters) are also easily formulated in food products and can be useful in functionalfoods. Furthermore, work on new formulations and crystalline forms of free phytosterols is nowbeing conducted with the aim of clarifying the true utility of these simpler and less expensivecompounds for cholesterol management. Such research efforts are beginning to answer some ofthe most intriguing questions regarding the biological activities and health-promoting effects ofthese phytochemicals. For instance, are phytostanols really more effective at lowering serumcholesterol levels than phytosterols? Which provide greater reduction in serum cholesterol levels,phytostanyl esters or phytosteryl esters? Are esterified phytosterols and -stanols more effectivethan free phytosterols/stanols in lowering serum cholesterol levels? What are the negative con-sequences of consumption of phytosterols and -stanols on fat soluble vitamin and antioxidantabsorption? Does consumption of phytosterols and phytostanols lead to elevated (or depressed?)serum levels of those species, and if so, what are the consequences of this on human health? Theanswers to these questions, surprisingly, have changed considerably over the last 10 years. Thecurrent perspectives on these issues and additional questions will be discussed in the sections thatfollow. While this review was being prepared, another review was published [150] and it providestimely insights into several of these questions.


4.2. Active forms and mechanism of action of steryl and stanyl esters

Phytosterols and phytostanols (primarily 4-desmethyl sterols and stanols) have been shown toinhibit the uptake of both dietary and endogenously-produced (biliary) cholesterol from theintestine. This results in a decrease in serum total and low-density lipoprotein cholesterol (LDL-C) levels [151], even for those individuals who are already on a low cholesterol diet [152,153].Levels of protective, high-density lipoprotein cholesterol (HDL-C) are typically not decreased bydietary phytosterol consumption. The exact mechanism by which phytosterols decrease serumcholesterol levels is not completely understood but several theories have been proposed [3,154].One theory suggests that cholesterol in the intestine, already marginally soluble, is precipitatedinto a non-absorbable state in the presence of added phytosterols and stanols. A second theory isbased upon the fact that cholesterol must enter bile-salt and phospholipid-containing ‘‘mixedmicelles’’ in order to pass through intestinal cells and to be absorbed into the bloodstream. Choles-terol is only marginally soluble in these micelles and it is displaced by phytosterols (and stanols),preventing its absorption. Unlike cholesterol, phytosterols and to a greater extent, phytostanols arevery poorly absorbed [155] and the small amount that is absorbed is actively re-excreted in bile. Thisresults in low serum levels of these sterol molecules.Although it has been the phytostanyl and phytosteryl fatty acid esters in margarine-like spreads

that have had recent commercial success in LDL-C-lowering efficacy, the physiologically activeforms are most likely the free, unesterified phytosterols and stanols. This is because steryl andstanyl esters are thought to be rapidly hydrolyzed by intestinal enzymes, yielding the physiologi-cally-active free phytosterol and phytostanol forms [3]. A recent in vivo study [156] confirmedthat in fact about 90% of ingested stanyl esters were hydrolyzed in the small intestine.

4.3. Recent clinical studies on phytostanyl esters

More clinical studies have been carried out to date on the effects of stanyl esters on blood lipidsthan on any class of phytosterols. These studies, which were recently reviewed [157], havedemonstrated the ability of stanyl esters to lower ‘‘bad’’ LDL-C levels by as much as 10–14% innormo- and hypercholesteromic adult males and females (with or without statin drugs), andchildren. Levels of ‘‘healthy’’ HDL-C and triacylglycerols were unaffected. A recent clinical study[158] confirmed that patients already taking statin drugs could reduce their LDL-C levels anadditional �10% by inclusion of stanyl ester margarines in their diets. Many studies havedemonstrated the efficacy of stanyl ester-enriched high-fat spreads in reducing LDL-C levels inindividuals consuming diets high in fat and cholesterol. Recently it was shown that stanyl estersare also effective in low-fat spreads fed to individuals on low-fat, low cholesterol (NCEP Step II-type) diets [153].Stanyl esters can be derived from tall oil, a phytosterol-rich byproduct from the pulping of pine

and other trees [148]. In that process, tall oil phytosterols are refined and purified, chemicallyhydrogenated to phytostanols, and then esterified with food-grade fatty acids. A similar productcan be produced starting with phytosterols derived from vegetable oil refining. Stanols derived viahydrogenation from tall oil contain primarily sitostanol and campestanol in a ratio of about 92:8.Stanols derived from soy oil contain a sitostanol/campestanol ratio of about 68:32. The twoblends appear to have equal efficacy [159] for lowering LDL-C levels.


4.4. Recent clinical studies on phytosteryl esters

Steryl esters are made by esterifying phytosterols derived from vegetable oils with food-gradefatty acids. Steryl esters are simpler and cheaper to synthesize than stanyl esters because nohydrogenation is required. The phytosterol composition of steryl esters depends upon their vege-table oil source and is frequently a mixture of sitosterol, campesterol, stigmasterol, brassicasterol,isofucosterol (�5-avenasterol), and other minor constituents.Like stanyl esters, phytosteryl esters (steryl esters) have been shown to consistently lower serum

LDL-C levels in clinical studies. Studies conducted up to the year 2000 have been recentlyreviewed [157]. Similar to the effect seen with stanyl esters, consumption of steryl esters resulted inLDL-C reductions up to �10%ormore, with no change seen inHDL-C values.Most of the clinicalstudies done prior to 2001 were conducted on normo- or mildly hypercholesterolemic individualsconsuming a high-fat steryl ester-enriched spread along with a high fat diet. More recent studies haveshown steryl esters to effectively lower LDL-C levels when administered in low-fat spreads and saladdressings [152,160] and when administered in various forms to individuals on low fat/low cholesteroldiets [152,160,161]. Individuals with different types of hypercholesterolemia [162] responded similarly(10–15% reduction in LDL-C) to steryl ester spreads and steryl esters were found to further reduceLDL-C levels in patients already being treated with hypocholesterolemic drugs such as statins [162]and fibrates [161]. A very recent study [163] reported that people with mild to moderatehypercholesterolemia all responded to a 1.8 g/day dosage of phytosteryl esters but the reductionin LDL-C levels were most marked in those with a high dietary intake of cholesterol, energy, totalfat, saturated fatty acids, and a high baseline cholesterol absorption.

4.5. Recent clinical studies on free phytosterols and phytostanols

Research on the cholesterol-lowering effects of free phytosterols and phytostanols has yieldedpositive but, inconsistent results due, at least in part, to the difficulty in formulating and deliver-ing these relatively insoluble substances. Heinemann and colleagues [164] showed that low doses(1.5 g/day) of free sitostanol was effective in lowering LDL-C levels. Becker and colleagues [165]reported that LDL-C levels in hypercholesterolemic children could be reduced by a surprising33% with only 1.5 g/day sitostanol, whereas administration of as much as 6 g/day of sitosterolresulted in a smaller (20%) reduction. Miettienen and Vanhanen [166] however, reported littleeffect from feeding subjects 0.7 g/day free sitostanol and Denke [167] reported no reduction ofLDL-C in patients given 3 g/day sitostanol (given in gelatin capsules rather than mixed into adietary component) along with a low-fat (NCEP step I) diet. More recently, Jones and colleagues[168] reported that a 1.7 g/day dose of tall oil phytosterols containing 20% sitostanol and 80%other phytosterols (primarily sitosterol and campesterol), reduced LDL-C 15% in hypercholes-terolemic men. Miettienen and Vanhanen [166], Pelletier et al. [169] and Sierksma et al. [170]showed that a dosage of 0.7–0.8 g/day of free sterols resulted in a LDL-C reduction of 6–15%.Most recently, a clinical study [171] showed that 1.8 g/day of free sterols, free stanols, or a freestanol/sterol mixture all gave statistically similar reductions in LDL-C in the range of 10–15%.Hence the preponderance of data indicates that free phytosterols and phytostanols are effective inlowering LDL-C, as long as they are formulated and delivered in a ‘‘bioavailable’’ physical state.Christiansen and colleagues [172] also recently reported that the effect of plant sterols is highly


dependent on their physical state and that hypercholesterolemic subjects consuming 1.5 or 3.0 g/day of free plant sterols in a ‘‘microcrystalline’’ (unesterified) form experienced a 7.5–11.6%reduction in LDL-C levels.

4.6. Relative LDL-C lowering efficacy: sterols versus stanols & esters versus free forms

Until about 1998, many clinicians and researchers and at least one manufacturer, believed thatstanyl esters were more potent LDL-C lowering agents than steryl esters and that stanols, ingeneral, were more effective than sterols at lowering serum LDL-C levels in humans. The ratio-nale for this assumption has been reviewed [3,173,174] and is also discussed in Section 4.5. Insummary, this conclusion was based on research in the 1950s, which indicated that up to 25 g offree sitosterol was required for maximum LDL-C reduction, and from the results of later studieswith both animals [175–177] and humans [164,165,174,178]. When combined, these studies indi-cated much greater cholesterol-lowering effects from stanols and stanyl esters. Because of thisevidence, considerable effort and expense were invested in development of stanyl ester products.Many of the studies cited above, however, were not done with systematic cross-comparison ofsterols and stanols in the same subjects and researchers do not always use the same methods tocompare and calculate the efficacy of their products in human trials.More recently, five head-to-head comparisons of stanyl esters versus steryl esters have been

completed [179–183]. Weststrate and Meijer [179] first compared the effects of margarines enri-ched with either vegetable steryl esters or sitostanyl esters (at 1.5–3.3 g/day) in mildly hypercho-lesterolemic patients. The two esters were equally effective in lowering blood levels of total serumcholesterol and LDL-C. A similar study confirmed these results with a low-fat diet regimen [180].Jones et al. [181] then reported that steryl esters were slightly more effective than stanyl esters inreducing total serum cholesterol and LDL-C levels with dosages of 1.84 g/day, in 15 hypercho-lesterolemic patients on a controlled diet. A unique recent study [182] with ileostomy patientsshowed that steryl and stanyl esters were equally effective at inhibiting the absorption of choles-terol from the intestine. Finally, another direct comparison of steryl and stanyl esters [183] notonly indicated equivalent efficacy in LDL-C lowering, but also on their effects on carotenoidabsorption. Based on the results [179–183] and analysis [184–186] of these studies, stanyl andsteryl esters appear to be roughly equivalent in their cholesterol-lowering efficacy.In light of the information discussed in Section 4.5, including the recent research of Jones et al.

[168] and others [171,172], it may be argued now that properly formulated free phytosterols andstanols may be as effective as stanyl and steryl esters for lowering LDL-C levels in humans. Thisseems to be confirmed in a recent study by Nestel et al. [187], who compared free stanols andphytosteryl esters head-to-head. The stanols and steryl esters were consumed (�2.4 g/day) incereal, bread and margarine. Sterol esters gave a 13% reduction in LDL-C while stanols gave an8% reduction. The researchers said the difference between the two values was not significant.While more research remains to be done to completely resolve this question, a current analysis ofrecent research appears to indicate that free sterols and stanols and steryl and stanyl esters, for-mulated in a manner that makes them ‘‘equally bioavailable,’’ all may give similar effects onLDL-C levels [188]. Because of the importance of this point to consumers, clinicians, manu-facturers, and researchers, this point will likely be studied and clarified by additional long-term(more than 3–4 weeks) head-to-head comparisons of all these different phytosterol forms.


4.7. Health claims

In September 2000, the FDA issued a rare interim final rule, allowing a health claim for reducingthe risk of coronary heart disease for foods containing plant stanyl and steryl esters [157] as long asthe foods were low in saturated fat and low in cholesterol. This was only the 12th time the FDA hasallowed a health claim. The Rule included stanyl esters in spreads, salad dressings, snack bars, anddietary supplements (soft gels), whereas it included steryl esters in only two of these applications,spreads and salad dressings. To qualify for the health claim, a product had to contain no more than13 g of total fat per serving or per 50 g. Spreads and salad dressings, however, were exempted fromthis requirement. Foods had to contain at least 0.65 g of phytosteryl ester or 1.7 g of phytostanylester per serving and at least two servings had to be eaten at different times of the day for a totalconsumption of 1.3 and 3.4 g/day of steryl and stanyl esters, respectively. The requirement formore than twice the amount of stanyl esters than steryl esters seems contradictory to the pre-ponderance of scientific literature that indicates similar efficacy for both chemical species. How-ever, the ruling was based upon the data available at that time and the fact than many of thestanyl ester studies were performed using relatively high dosages. The FDA allowed two separatecomment periods with a final rule expected sometime in 2002. Comments sent to FDA focused onissues surrounding dosage levels for steryl and stanyl esters, whether free phytosterols and phy-tostanols should be included in the rule, additional food applications, and whether consumptionof these ingredients may have a negative effect on absorption of dietary carotenoids. A review ofthese and other issues surrounding the health claim was recently published [189].

4.8. Reduction in the risk of coronary heart disease

The FDA Interim Final Rule gives examples of wording that can be used on labeling of phy-tosterol-containing products. Generally, it may be said that consumption of these phytonutrientsin the appropriate amounts ‘‘may’’ decrease the risk of heart disease. Estimates for the actualamount of risk reduction that could be expected from consuming phytosterol-containing pro-ducts will probably not be part of the Health Claim but this point was addressed [179]. Theseresearchers estimated that consumption of 3 g/day of stanol or sterols could reduce the risk of heartdisease by 15–40% depending upon age and other dietary factors. Another recent review [184]indicated that consumption of 2 g/day of sterols or stanols could result in a reduction in the risk ofheart disease by about 25%. This effect was considered to be more significant than reducing theintake of saturated fat.

4.9. Toxicology, anticancer properties and potential benefits

It is estimated that well over 2400 subjects have taken part in clinical studies with phytosterolsand stanols with dosages up to 25 g or more per day and no adverse effects were noted. The drugCytellin (primarily sitosterol) was prescribed for more than 20 years and had an excellent safetyrecord. Concern had been expressed (reviewed in [3]) that ingestion of large amounts of phytos-terols might lead to high serum levels and consequent estrogenic or atherogenic effects. This isone reason that much work on commercial products in the early 1990s was focused on the largelyunabsorbed stanols and stanyl esters. A series of studies (reviewed in [185]) conducted over the


last 4 years have indicated a complete lack of toxicity in animal models and humans, except forindividuals with the extremely rare autosomal recessive disorder called phytosterolaemia (sitos-terolaemia) [190,191], and these atypical patients are strongly urged to avoid dietary intake ofphytosterols. Some research studies [164,165] however, suggested that consumption of sitostanolby patients with sitosterolemia could be an effective way to reduce their levels of serum choles-terol and free phytosterols, which have recently [192] been shown to occur in oxidized form inpatients with this condition. These oxidized forms are analogs to the oxidized forms of cholesterolwhich are thought to be atherogenic and cytotoxic. At present, it is not known if these oxidizedforms of phytosterols have negative effects on human health and in view of all the safety dataavailable, stanyl and steryl esters are now listed as GRAS (Generally Recoginzed as Safe) by theFood and Drug Administration for the general population. Studies [193] conducted since thisGRAS certification confirm this view. Phytosteryl esters were well tolerated and caused noadverse effects at daily intakes up to 9 g/day during an 8-week period [193].Several studies have indicated that phytosterols may have health-promoting effects such as anti-

cancer activity. Awad and colleagues [194–196] provided evidence that phytosterols were toxic tobreast cancer cells. Other studies indicated that phytosterols were toxic to colon cancers [197,198].Other studies indicate that phytosterols may be one of the active ingredients in saw palmettowhich contributes to its toxicity towards prostate cancer cells [199]. If these results are confirmed,then the presence of some reasonable levels of serum phytosterols may be desired and conversely,consumption of any dietary component that would lower those levels may be detrimental to ourhealth. In this regard, a review of the literature would indicate that consumption of free sterols,free stanols, and their esterified forms all may have a differing effect on serum phytosterol levels.As noted in clinical studies [179,180] and in a recent review [3], consumption of largely unab-sorbable stanols and stanyl esters results in a lowering of all types of serum phytosterols. On theother hand, consumption of phytosteryl esters leads to increases in blood phytosterol levels [179].For instance, administration of 2.5 g/day of plant steryl esters resulted in an approximate dou-bling of the subjects’ total serum phytosterol levels [162]. While steryl esters raise serum phytos-terol levels and stanol decrease them, two studies have suggested that consumption of freephytosterols and mixtures of free phytosterols and phytostanols may result in no change (neitherincrease or decrease) in serum phytosterol levels [168,200]. Additional research will be needed toconfirm this point.There is evidence that some phytosterols may have antioxidant activity. White and Armstrong

[201] demonstrated that �5 avenasterol, which occurs in high levels in oats, may have valuableantioxidant activity. Additional studies have confirmed that �5 avenasterol and other phytoster-ols that contain an ethylidene group possess antioxidant and antipolymerization properties,especially valuable during frying [202–204]. Whether these antioxidant properties have any sig-nificance to human health remains to be seen.

4.10. Effects on absorption of fat soluble vitamins and antioxidants

Several studies [179,180,205–207] have shown that consumption of stanyl and steryl estersslightly reduces the absorption of the fat-soluble antioxidants b-carotene (pro-vitamin A), lyco-pene, and a-tocopherol (vitamin E). Law [184] reviewed this data and stated that plant sterols andstanols lower blood concentrations of b-carotene by about 25%, concentrations of a-carotene by


10%, and concentrations of vitamin E by 8%. However, a key role for these vitamins may be toprotect LDL-C from oxidation. Since sterols and stanols reduce the amount of LDL-C, andbecause lipophilic carotenoids and tocopherols are known to be associated with LDL particles, itmay be appropriate to adjust (or correct) the blood concentrations of these vitamins for the lowerLDL-C concentrations. With this adjustment, stanols and sterols did not significantly lowerblood concentration of vitamin E but concentrations of b-carotene were reduced by 8–19% orless [153]. In view of these results, Hendriks et al. [206] suggested limiting daily dosage of sterylesters to about 1.6 g/day, a dosage that gave good LDL-C reductions without seriously affectingplasma carotenoid concentrations. A later meta-analysis [188] questioned whether limiting thedose to 1.6 g would be necessary since the effects of steryl and stanyl esters on carotenoids andtocopherols were not necessarily dose dependent. More recent data [152] has confirmed the factthat stanyl and steryl esters do affect serum carotenoid levels and this finding has led to sugges-tions that patients consuming ester-containing spreads should either increase their intake of car-otenoid-rich foods like apricots, cantaloupe, broccoli, and spinach [152] or take supplementscontaining these carotenoids to offset the decrease in absorption [162]. This has now beenattempted in one clinical study which indicated that an increase in dietary carotenoids whenconsuming plant sterols or stanols was effective in maintaining plasma carotenoid levels [183].A final point regarding fat soluble vitamins that may be of importance was noted in a recent

review [186]. The review noted that administration of free phytosterols and phytostanols may notinduce maladsorption of fat soluble vitamins and antioxidants as much as that caused fromconsumption of the fatty acid ester forms. If this is verified in more studies, it might bring evenmore attention to use of the free phytosterols and phytostanols in functional foods.

4.11. Dosage levels and frequency

There have been some differences in opinion regarding the recommended dosages for phytos-terols and phytostanols. The FDA’s interim final rule suggested that more than twice the amountof stanyl esters (3.4 g/day) than steryl esters (1.3 g/day) were needed to ensure a significantreduction in LDL-cholesterol levels. As stated in Section 4.6, it is now thought that effects ofthese esters are relatively equivalent. Studies indicate that there is a dose-dependent response onLDL-C lowering for both steryl [206] and stanyl [208] esters. Because of these and other studies, arecent meta-analysis [184] suggested that a dose of 2 g/day of either steryl or stanyl esters shouldgive optimum effect and this 2 g/day dosage has now been recommended in the latest publishedNational Cholesterol Education Program guidelines [209].Dosage frequency is another area that is being re-evaluated. It had been commonly believed

that maximum LDL-C lowering efficacy of plant sterols and stanols is contingent on them beingconsumed along with every meal. The FDA’s Interim Final Rule for the Health Claim requiresconsumption of steryl and stanyl esters in at least two servings per day. However, it has beenpointed out [184] that LDL-C lowering effects were similar if stanyl and steryl esters were con-sumed with either 2 or 3 meals per day. Another study [210] indicates that the cholesterol loweringeffect of plant stanols appears to be independent of whether they are consumed once per day ordivided over three meals. This finding is very significant from at least two points of view. First,most Western consumers’ lifestyles do not permit the incorporation of phytosterols into foods intwo or three meals per day. Being able to benefit from consumption of a phytosterol-enriched


food at only one meal greatly increases the chances of consumers ‘‘buying in’’ to the use of thesefunctional foods. The other significant aspect of these findings is that it means that the mechan-ism by which sterols and stanols work may need to be re-examined. The study suggests that plantstanols either remain in the intestinal lumen or possibly are associated with enterocytes [210].Further, they hypothesized that plant stanols have effects on lipoprotein metabolism beyond theireffect on the micellar solubility of cholesterol. These unexpected findings indicate that moreresearch should be done to fully understand this phenomenon.

4.12. Phytosterol/phytostanol products: past, present, and in development

Stanyl ester products entered the market in Finland in the mid 1990s, followed by both stanyland steryl ester products in other European countries (Table 4). Stanyl and steryl ester productsentered the US market in 1999 after considerable regulatory discussions and delay. McNeilConsumer Healthcare (with North American marketing rights to Raisio’s Benecol) intended tolaunch Benecol in the fall of 1998 as a dietary supplement which, under the DSHEA act is exemptfrom many US FDA regulations. The FDA, however, considered Benecol to be a food or foodingredient. McNeil agreed to market Benecol as a food, but that required either a new foodadditive petition, or recognition of GRAS status. In 1999, a panel of independent experts in theUSA concluded that plant steryl esters were GRAS for use as an ingredient in vegetable oil-basedspreads in amounts not to exceed 20%. Based upon this GRAS recognition, the FDA approvedspreads containing up to 20% of plant steryl and stanyl esters. Benecol (containing stanyl esters)was then finally launched in May of 1999. Also in 1999, Lipton, and its parent company, Uni-lever, launched their product, ‘‘Take Control,’’ which contained vegetable oil-derived sterylesters. Because Take Control’s active ingredients, steryl esters, were simpler and less expensive toprepare (no hydrogenation required), Lipton’s product could be sold at a lower price than Ben-ecol. This initially did not seem to be a major advantage since steryl esters were considered bymany to be less efficacious than stanyl esters. However, with the present evidence that steryl andstanyl esters are equivalent in reducing serum cholesterol levels, Take Control has a significantmarketing advantage over stanyl ester products in this regard.In October 2000, Forbes Medi-Tech Inc. announced the first consumer product test market of its

phytosterol-based cholesterol lowering ingredient, Phytrol (unesterified tall oil phytosterols) in Aus-tralia and the USA. Phytrol is exclusively licensed worldwide to Novartis Consumer Health, Inc. andin 2000, Novartis announced a joint venture with Quaker Oats to form Altus Foods, which willmanufacture ‘‘healthy foods’’ containing Phytrol. The successful marketing of a free phytosterolproduct would be quite an accomplishment in view of previously published evidence that free sterolsare difficult to formulate and that their poor bioavailability necessitates large dosages for significantreductions in serum LDL-C levels. However, in view of the most up-to-date clinical informationdiscussed in Sections 4.5 and 4.6, it would seem reasonable to assume that such products are not onlypossible, but they might have a marketing edge, since the lower cost of free, nonesterified and non-hydrogenated phytosterols could result in a less expensive product. Forbes also claims that Phy-trol (consumer branded as ‘‘Reducol’’) can be used in a wide variety of food products besidesfatty foods. If this is confirmed, it could also help marketing efforts for Phytrol.A unique oil was discovered in corn fiber (a fiber-rich byproduct from corn wet-milling) by a

team of scientists with USDA’s Agricultural Research Service (ARS) [51] and proposed as a


Table 4Major phytosterol/phytostanol products being marketed in March 2002

Product trade names Phytosterol activecomponent(s)

Type of food Manufacturer Website Marketing status

Benecol Phytostanyl esters Margarine Spreads

(US)

Raissio/McNeil www.benecol.com Presently available

US and EuropeMargarine spreads,cream cheese spreads,

milk, Mayonnaise,yogurt, Meat products,snack bars (Europe)

www.benecol.co.uk

Take Control (US) Phytotosteryl esters Margarine spreads,salad dressings

Lipton/Unilever www.takecontrol.com Presently availableUS and International

Flora Pro-activ and

Becel Pro-activ(Europe and Canada)

www.floraproactiv.co.uk www.becel.

com.br/becelproactiv

Phytrol Free phytosterols Various Forbes Meditech www.forbesmedi-tech.com/news2001/072401.asp

Presently availableas an ingredient

Reducol Phytrol Various Novartis www.consumer-health.novartis.com/vmsi.html

Presently availableas an ingredient

Cholesterol

Success

Reducol Tablets Twinlabs Co. www.twinlab.com/prod.cfm?product

=reducol

Unknown

Diminicol Phytosterols Unknown Teriaka www.teriaka.com In developmentCholestatin Free phytosterols Tablet Degussa

Bioactives

www.bioactives.de/bioactives/html/e/

products/factsheets/factsheets.htm

Presently available

Prosterol Free phytosterolsand policosanols

Tablet In Development

Logicol Phytosterols Spreads and Milk Meadow Lea http://www.logicol.com.au Spreads now

available AustraliaChoLESStolife Phytosterols and

phytostanolsPowder Lifeline

Techolgieshttp://www.lifelinetechnologies.com/products.html

In Development

CardioAid-L Phytosterols andlecithin

Beverageingredient

ADM http://www.admworld.com/try/tryadmproducts.htm

In Development

CardioAid-P Phytosterols and

soy proteins

492

R.A.Moreauetal./Progress

inLipidResea

rch41(2002)457–500

natural cholesterol-lowering oil. Corn fiber oil, called Amaizing Oil, was shown to lower serumcholesterol levels in an animal model at the University of Massachusetts (UMass) and was thenpatented [211] as a joint invention of ARS/UMass. Additional animal studies [212,213] have sinceconfirmed its cholesterol lowering effects. Corn fiber oil has both phytosterol and phytostanylesters and hydroxycinnamate esters (Fig. 7) and the latter may impart functional properties notfound in any current commercial phytosterol product. Unlike phytosterols in soy or tall oil, mostof the phytosterols in corn fiber oil are naturally esterified with either fatty acids or phenolicacids, such as ferulic acid, a potent antioxidant [211]. Furthermore, corn fiber oil contains a highlevel of sitostanol in the ferulic acid ester fraction. In fact, corn fiber oil appears to be the richestsource of natural stanols (and stanyl esters) ever reported. Corn fiber oil also contains g-toco-pherol and various carotenoids, both with important antioxidant properties. The levels of totalphytosterols in corn fiber oil range from about 15 to over 50%, depending on extraction and fiberpre-treatment conditions [51]. Levels of g-tocopherol also vary with fiber pre-treatment condi-tions [214] and range from about 0.3 to 3%. Corn fiber oil’s combination of natural cholesterollowering components and antioxidants, which could potentially prevent oxidation of LDL-cho-lesterol, could give a ‘‘one-two punch’’ in the fight against heart disease. The ARS scientists haveworked with several major US corporations to move the oil towards commercial products, whichcould include spreads, chocolates, dairy products, cooking oils, and other foodstuffs with health-promoting properties. An exclusive license for manufacturing and use of the oil for non-cookingoil applications has been granted. The use of the oil in cooking oil applications is available forlicensing.During the years 2000 and 2001, Procter & Gamble (P&G) test marketed a new line of phy-

tosterol-containing cooking oils under the brand name, ‘‘CookSmart’’. These oils contained soyphytosteryl esters. P&G was the first company to market a phytosterol-containing cooking oilwhich conceivably could add cholesterol-lowering phytosterols to fried foods such as french fries.Forbes Medi-Tech Inc. announced in December 2000 its development of a ‘‘designer oil’’ which

reduces LDL-C and increases energy expenditure, and hence may prevent people from gainingweight. The research was conducted at McGill University between October 1999 and May 2000.The oil contains Forbes’ phytosterol based ingredient, Phytrol, which is incorporated into the oilby a proprietary process that preserves clarity of the oil. Another type of phytosterol productattracting attention is a product that contains phytosterols in a diacylglyerol (versus triacylgly-cerol) base [215]. Another study [216] demonstrated that the effect of only 500 mg/day of phy-tosterols in this product reduced serum LDL-C levels by about 8% compared to the sameamount of phytosterols in a triacylglycerol base, which caused no decrease in LDL-C levels.Some consumer resistance to current commercial products stems from the high fat content of

the spreads and margarines in which they are formulated. Recent research shows that phytoster-ols delivered in a low fat spread are as effective as those in higher fat formulations (see Sections4.3 and 4.4). Low fat spreads containing phytosterols are now commercially available.Another approach to delivering phytosterols in a non-fat form is taken by Degussa Bioactives

with their commercial product, Cholestatin. Cholestatin contains a mixture of free phytosterols ina capsular form containing no added fat.There is a demand for phytosterol formulations that could be included in beverages, dairy

drinks, and non-fat foods. ADM has developed a patent-pending phytosterol formulation thatallows introduction of sterols in a dispersible form in an aqueous application [217].


Monsanto recently received a patent on a ‘‘Phytosterol protein complex’’ which is com-prised of phytosterols, proteins and edible oil [218]. The complex is said to ‘‘increase thebioavailability of phytosterols’’ and that ‘‘It is most preferred to extract the phytosterolsfrom corn fiber oil’’.Lecithin also plays a valuable role in increasing the bioavailability of free phytosterols. A recent

study [219] showed that sitostanol powder (1 g) reduced cholesterol absorption in human subjectsby about 11%. In contrast, only 300 mg of sitostanol administered in lecithin micelles wererequired to reduce cholesterol absorption by 34%.More than 20 US patents on phytosterol and phytostanol products or processes have issued in

the last 10 years (most of these US patents were filed as part of a ‘‘family’’ of internationalpatents that can be accessed at www.uspto.gov). These patents fall into several categories. Theearliest and largest group are patents on phytostanyl esters filed by Raisio and McNeil [148,220–226]. Lipton (Unilever) has several patents on phytosterol/phytostanol technology [227–230].Other major patents include various one from Forbes Medi-Tech [231]; a lecithin–phytosterolpreparation [232,233] owned by Washington University; another phytosterol–protein complexowned by Monsanto [218]; a phytosterol–emulsifier complex owned by Kraft [234,235]; and a‘‘nanoscale’’ process to produce emulsions containing microscopic particles of phytosterols [236].Finally, several patents involve multi-component phytosterols products include: our own patenton the extraction, composition of mater and cholesterol-lowering applications of ‘‘corn fiber oil.’’(which contains natural steryl esters, free sterols, hydroxycinnamate steryl and stanyl esters,tocopherols, and carotenoids) [211]; a multi-component formulation that combines tocopherolsand tocotrienols (10–30%), free sterols (2–20%), steryl esters (2–20%), cycloartenols (0.1–1.0%)and saturated fats (7–19%) [237], and a formulation that contains phytosterols (1.2–20 wt.%),dissolved in diacylglycerol (15 wt.% or more) [215].

5. Conclusions

Phytosterols represent a diverse group of natural products, and knowledge about their occur-rence in various plants and their chemical composition has gradually accumulated during the last100 years. Much early research and utilization of phytosterols focused on their value as pre-cursors in the synthetic synthesis of several steroid hormones. During the last 10 years, there hasbeen an unprecedented escalation of interest in phytosterols. Most of this interest has focused onthe cholesterol-lowering properties of 4-desmethyl phytosterols and phytostanols, and evidence ofthis phenomenon include at least 25 clinical studies on phytosterol products, 20 patents on phy-tosterol products, and at least 10 major commercial phytosterol products currently being mar-keted in many parts of the world.

Acknowledgements

The authors would like to thank the following persons for generously providing valuableinformation and insights for this review: E. Harrison, A. Lichtenstein, G. Meier, G. Picchinoni,V. Pirronen, I. Wester, and J. Zawistowski.


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