ReviewEnzymes as biocatalysts in the modification ofnatural lipidsFrank D Gunstone*Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, Scotland

Abstract: Though designed by nature to effect hydrolysis of lipids, lipases can, under appropriate

reaction conditions, promote ester formation through reaction of acids and alcohols (esteri®cation) or

of esters with acids (acidolysis), alcohols (alcoholysis), or other esters (interesteri®cation). Compared

with chemical processes already carried out on an industrial scale enzymic reactions occur under

milder (and `greener') conditions though they may take longer. Of greater signi®cance is the speci®city

shown by the enzymes which permits the formation of lipid derivatives not easily prepared by

conventional laboratory procedures.

This review describes the lipases and their various speci®cities and reports on their use in hydrolysis

and in the production of phospholipids, fatty acids, alkyl esters, mono- and di-acylglycerols, tri-

acylglycerols, other esters, and amides. Some of these have already led to marketable products but for

the most part the full potential of these reactions has yet to be realised. The reactions of other enzymes

promoting interesting reactions at unsaturated centres are also described.

# 1999 Society of Chemical Industry

Keywords: lipases; enzyme-catalysed hydrolysis; esteri®cation; acidolysis; alcoholysis; interesteri®cation;polyunsaturated fatty acids; medium-chain triacylglycerols; mono-, di- and tri-acylglycerols; wax esters; sugaresters; amides; enzyme-catalysed reactions; ole®nic centres

1 INTRODUCTIONCommercial oils and fats are now produced at levels

around 100 million tonnes per annum and are used

mainly for food (80%), animal feed (6%), and for

oleochemical purposes (14%) the great majority

(90%) of which is for the production of soaps and

other surface-active materials. On this basis it is not

surprising that much research in the lipid ®eld is

devoted to nutrition which also drives many of the

processing techniques. The natural materials are not

always best suited for the nutritional demand made on

them and for the whole of the 20th-century (and

before) lipid scientists and technologists have been

engaged in improving quality in nutritional terms by

blending, fractionation, partial hydrogenation, and

interesteri®cation. Some of these developments have

brought their own problems as in the recognition that

acids with trans con®guration, formed during partial

hydrogenation, as well as saturated acids are not

nutritionally desirable beyond very modest levels.

During this century partial hydrogenation, fractiona-

tion, and interesteri®cation have been developed and

used extensively to modify lipids. Despite, or because

of, their failings these processes are being improved

continually by the makers of equipment and catalysts.

Other drivers come from the growing demand for

`natural' foods and from environmental concerns to

reduce energy requirements and to minimise waste as

well as economic pressures to produce good products

at acceptable prices.

Another area of R and D is the production of `better'

natural products, ie lipids with a more desirable fatty

acid and triacylglycerol composition. These are being

developed in a number of ways which were described

in a recent review.1

. The domestication of wild plants (eg cuphea,

calendula, etc) often to produce high levels of a

single acid for oleochemical purposes.. Modi®cation of existing oilseed plants by con-

ventional seed breeding as in the development of

double zero rape seed from high-erucic rape seed

oil or by genetic modi®cation. So far the latter

have been directed mainly to the production of

species which show agricultural traits such as

herbicide resistance and pesticide resistance, etc.

Journal of the Science of Food and Agriculture J Sci Food Agric 79:1535±1549 (1999)

* Correspondence to: Frank D Gunstone, Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, ScotlandE-mail: fgunst@scri.sari.ac.uk(Received 1 March 1999; accepted 18 May 1999)

# 1999 Society of Chemical Industry. J Sci Food Agric 0022±5142/99/$17.50 1535

Only recently have materials with changed fatty

acid composition begun to appear. The ®rst

example is a high-laurate canola but other

interesting materials are following quickly.. Production of oils from microbial sources such as

oils rich in arachidonic acid (Mortierella alpina)and in docosahexaenoic acid (Crypthecodiniumcohnii).

This review is devoted to a different area of lipid

research viz the use of biocatalysts in general and

lipases in particular to effect changes in the lipid

mixtures produced naturally. The lipases have long

been known as important enzymes in lipid metab-

olism, promoting complete or partial hydrolysis of

glycerol esters. Attempts have been made to exploit

these as hydrolysis catalysts and the following quota-

tion is taken from a book published in 19272 at the end

of a two-page description of technical hydrolysis with a

preparation from castor seeds. `The lipase fat-splitting

process is a step in the right direction in that it is an

attempt to use Nature's active chemical agents ± the

enzymes which by operating at natural temperatures

avoid the unnecessary expenses of fuel for steam and of

manufactured chemicals as reagents'. Those prescient

word were penned three-quarters of a century ago. In

the intervening years steps in this direction have been

slow and faltering but now they are gathering pace. In

the last ten years lipase-catalysed reactions have been a

subject of increasing interest and growing importance

resulting from the realisation that, under appropriate

conditions, the biocatalysts can promote ester forma-

tion as well as ester hydrolysis and that, with careful

choice of lipase, reaction conditions and substrates, it

is possible to control acylation and deacylation to

produce speci®c fatty acids and triacylglycerols. This

ability has developed at the same time as a demand for

speci®cally structured molecules not readily available

from natural sources.

There is much research activity in this ®eld. Con-

ferences on lipase reactions are frequent and popular.

Many interesting molecules and mixtures have been

produced but only a few have led to commercial

products.1 The increased cost of using enzymes means

that so far only low-volume, high-value products have

been produced in this way. Nevertheless the wide

interest in this topic, in both academic and industrial

laboratories, suggests that further products will soon

come to the marketplace.

2 LIPASESCommercial lipases are available from animal, plant,

and microbial sources including those listed in Table

1.

Though designed for hydrolysis of lipids and related

Table 1. Selectivity shown by lipases fromselected sources

Lipase source

Glycerol

reactivity Discrimination against

Reaction

con®ned to

Aspergillus niger 1,3 �2

Aspergillus sp None

Candida rugosaa None D4, 5, and 6 acids/esters

C antarcticab None D4, 5, and 6 acids/esters

C lypolytica 1,3>2

C parapsilosis

Chromobacterium viscosum

Geotrichum candidum D9c acids

Humicola lanuginosa 1,3 �2

Mucor javanicus 1,3>2

Mucor mieheic 1,3>2 D4, 5, and 6 acids/esters

Oats (Avena sativa) D9 acids

Pancreatic (porcine) 1,3>2

Pre-gastric esterase 1,3>2

Penicillium sp 1,3>2

Phycomyces nites

Pseudomonas cepacia

Pseudomonas ¯uorescens None

Pseudomonas spp

Rape seedlings

Rhizomucor mieheic D4, 5, and 6 acids/esters

Rhizopus arrhizus 1,3>2

Rhizopus delemar 1,3 �2 D4, 5, and 6 acids/esters

Rhizopus japonica

Rhizopus javanicus 1,3>2

a Previously known as Candida cylindracea.b Also referred to as Novozyme 435TM.c Alternative names for the same lipase±also referred to as LypozymeTM.

1536 J Sci Food Agric 79:1535±1549 (1999)

FD Gunstone

molecules, lipases can also catalyse other reactions

associated with acyl groups. These include:

. Hydrolysis: reaction of ester with water producing

acid and alcohol and with hydrogen peroxide to

give peroxy acids.. Esteri®cation: the reverse of hydrolysis ± produc-

tion of ester from acid and alcohol.. Alcoholysis: reaction of an ester with a mono-

hydric alcohol such as ethanol, butanol, lauric

alcohol, or a polyhydric alcohol such as glycerol to

produce an ester with a different alkyl group.

Similar reaction with amines leads to amides.. Acidolysis: reaction of an ester with an acid

leading to exchange of acyl groups.. Interesteri®cation: reaction of one ester with

another leading to scrambling (randomisation)

of acyl and alcohol moieties.

These processes can be selected by choosing appro-

priate substrates and reaction conditions. Increasing

numbers of research papers utilise lipases as bio-

catalysts and cover a range of reactions. Perhaps the

most signi®cant are those aiming to produce triacyl-

glycerols of de®ned composition, mainly on the basis

of their nutritional properties (see Section 8).

Lipase-catalysed processes have attracted attention

because of the mild reaction conditions under which

they occur and the selectivity displayed by these

catalysts. In both these respects they differ from

typical chemical reactions.

Because enzymic reactions occur under mild con-

ditions of temperature and pH and at normal pressure,

they generally require less energy and are conducted in

equipment of lower capital cost than many other

chemical processes. Also, under these milder condi-

tions, the products are purer and less degraded

through alternative high-temperature reactions, so

they are more easily puri®ed and waste disposal is less

of a problem.

Because of the selectivity of many lipases it is

possible to obtain products which are dif®cult to make

by more conventional chemical reactions. These

enzymes may show one or more of several types of

selectivity (Table 1):

. Some distinguish between acids of different chain

length, reacting differently with short, medium

and long-chain acids.

. Reactivity is also in¯uenced by structural features

in the alkyl group of the fatty acid chain such as

double bond con®guration and position with

respect to the acyl function, and the presence

and position of other functional groups such as

branched methyl.. Reaction may differ with the nature of the acyl

source±whether it is free acid, alkyl ester, or

glycerol ester.. In esters of polyols (such as glycerol esters) there

may be a distinction between acyl chains linked to

primary and secondary hydroxyl functions (ie

those in the sn-1/3 and the sn-2 positions) and in

the relative reactivities of monoacylglycerols,

diacylglycerols, and triacylglycerols.

It is likely that speci®city will be further developed with

a fuller understanding of the optimum reaction

conditions (temperature, pH, molar ratio of reactants)

and with better enzymes resulting from modi®cation of

existing lipases or from new sources. This is an area of

active research.

The molecular basis for lipase activity and speci®city

is poorly understood but recent advances in this topic

have been reported for the lipases from Geotrichumcandidum and Candida rugosa,3 Pseudomonas cepacia,4

Rhizopus delemar,5 and for pancreatic lipase.6 It is

believed that lipase activity is associated with a serine

residue in the protein chain. Binding sites, associated

with speci®city, include a hydrophobic cleft or an

L-shaped hydrophobic tunnel approximately 20

carbon atoms long. Phycomyces nites after N-acylation

(C18±C22) of its lysine residue is 40 times more

reactive and has modi®ed selectivity in transesteri®ca-

tion reactions.7 Lipase activity can also be modi®ed by

the presence of surfactants as reported for Rhizopusjaponicus and sorbitan monostearate.8

To demonstrate the speci®city of the lipases from

Rhizopus delemar and Rhizomucor miehei, Sonnet and

Sazzillo9 examined the partial hydrolysis of the

triacylglycerol POS and compared the results with

those obtained with the pancreatic lipase used

analytically on the basis of its 1,3-speci®city (Table

2). Complete speci®city should lead to no oleic acid in

the liberated acids though this value must increase if

hydrolysis proceeds beyond 67%. Signi®cant selectiv-

ity of other kinds has also been reported for Geotrichumcandidum10,11 and for Penicillium sp.12 Many enzymes

Table 2. Demonstration of 1,3-selectivity inthe partial hydrolysis of POS

Glycerol esters Free acid

Lipase source Hydrolysis (%) P (%) O (%) S (%) P (%) O (%) S (%)

Porcine pancreas 39 27 48 24 43 13 44

Rhizopus delemar 51 21 60 18 45 3 52

Rhizomucor miehei 78 19 65 10 36 24 40

P=palmitic

O=Oleic

S=Stearic.

J Sci Food Agric 79:1535±1549 (1999) 1537

Enzymes as biocatalysts in the modi®cation of lipids

discriminate against acids with D4, D5, and D6

unsaturation as is fully demonstrated in Section 5.

Speci®city usually refers to hydrolysis of triacylgly-

cerols and is not always the same in esteri®cation

reactions. For example, Osterberg et al13 reported that

lipases from Rhizopus delemar and from other Rhizopussp were unable to promote hydrolysis of tri-g-linolenin

but promoted acidolysis of triolein by g-linolenic acid

when the latter was used in large excess.

New sources of lipase have been identi®ed in

germinating rape seedlings and in oats (Avenasativa).14±16 The latter displays a similar speci®city to

that shown by Geotrichum candidum except that, unlike

this lipase, it promotes reactions with elaidic acid and

its esters (9t-18:1). Hou et al17 and Hou18 have

reported a search for new lipases, several of which

show promise as useful catalysts.

Reaction systems usually consist of lipase (water-

soluble, though sometimes used in an immobilised

form) with its associated water, the reactants, perhaps

a solvent (water and/or organic solvent), and possibly

additional materials such as silica or a molecular sieve.

Reaction conditions must be controlled so that sub-

strates are present at the reactive interface in appro-

priate form and proportion and products are removed

from the interface so that they do not react further nor

inhibit the desired reaction. For example, silica gel is

often added to reactions involving glycerol since this

molecule is so strongly adsorbed on the lipase that

reaction is inhibited. Instead the glycerol is adsorbed

by the silica and made available to the enzyme only in

appropriately restricted amounts.19

Reactions promoted by lipases are generally rever-

sible, leading to an equilibrium. To get high yields of

the desired product it may be necessary to displace this

equilibrium by removing one of the products. Water

and other volatile products may be removed by

operating at a suitable temperature and pressure or

by using a molecular sieve as adsorbent. Other

products can sometimes be removed from the reaction

mixture by crystallisation and it is for this reason that

reaction may be started at one temperature (say 40 or

50°C) and then reduced to a lower temperature (say

10°C). If one product then crystallises from the

reaction mixture the equilibrium is displaced and

more of that product is formed.

3 HYDROLYSISFat splitting is normally effected by a continuous,

high-pressure, uncatalysed, counter-current process at

250°C and 20±60 bar. Under these high-temperature

conditions the products become discoloured and both

the fatty acids and the glycerol may have to be distilled.

Enzymic hydrolysis can be effected under milder

conditions with a wide range of lipases including

Rhizomucor miehei,20 Aspergillus sp,21 Pseudomonas¯uorescens,22 Candida rugosa,20 and Candida cylindra-cea23 acting at �37°C for about 24hours. High-

melting fats can be hydrolysed under these conditions

in the presence of a little iso-octane.24 Partial hydro-

lysis, exploiting the speci®city of lipases which

discriminate against particular acids to concentrate

these acids, has been discussed in Section 5.

4 PHOSPHOLIPIDSPhospholipases A±D provide for the speci®c hydrolysis

of each of the four ester bonds present in phospho-

lipids, and phospholipase D has been used for

transphosphatidylation, ie exchange of the alcohol

moiety attached to the phosphatidic acid residue.

Phospholipase A2 (or other lipases) can be used to

effect exchange of acyl groups.

Na et al25 used phospholipase A2 to prepare phos-

phatidylcholines enriched with n-3 PUFA. With this

enzyme phosphatidylcholine from egg yolk was hydro-

lysed to lysophosphatidylcholine (30°C, 16h) which

was then re-acylated (30°C, 16h) with a polyunsatu-

rated fatty acid (PUFA)-enriched mixture of acids

containing eicosapentaenoic acid (EPA) (41%) and

docosahexaenoic acid (DHA) (30%). The phospha-

tidylcholine ®nally contained 16 and 11% respectively

of these two acids. Using a 90% concentrate of DHA,

phosphatidylcholine with 35% of DHA was prepared.

Three commercially available lipases (Humicolalanuginosa, Rhizopus delemar, and Candida rugosa)

promoted hydrolysis of phosphatidylcholine to the

lyso derivative or to phosphatidic acid in solvents of

appropriate polarity.26 Egg yolk phospholipids

normally contain arachidonic acid (4.9%) and docosa-

hexaenoic acid (3.5%). Hara et al27 made a compara-

tive study of 19 commercially available lipases for the

hydrolysis of phosphatidylcholine to lyso compounds.

Highest reactivity was shown by lipases from Mucorand Rhizopus species. Virtually no hydrolysis occurred

with Candida lipases. When ®sh oil is incorporated into

the feed these values are changed to 1.5 and 10.9%

and after partial hydrolysis with Mucor miehei lipase

they are raised further to levels of 8.8 and 13.7%.28

Acidolysis of phosphatidylcholine with heptadecanoic

acid (Rhizopus arrhizus, 40°C) gave a product with

98% and 1% of this acid at the sn 1 and 2 positions

respectively indicating a successful and selective acid

exchange.29 Mustranta et al30 used Rhizopus mieheilipase (40°C, 24h) and oleic acid to convert di-

myristoylphosphatidylcholine to its 1-oleyl derivative.

Alcoholysis of phospholipids with ethanol, isopro-

panol or butanol under the in¯uence of lipases from

Mucor miehei or Humicola lanuginosa yields lysophos-

pholipids in high yield (22°C, 24h, 98%). By varying

the reaction time useful mixtures of phosphatidylcho-

line and lysophosphatidylcholine were obtained.31

The 1-lysophospholipid is converted to the 2-isomer

by reaction with ammonia vapour at 75°C. In the

presence of Mucor miehei lipase soy phospholipids

undergo alcoholysis with a range of alcohols (C4±C18)

at the sn-1 position (24h, 70±80% conversion) to give

lysophospholipids. There is no speci®city with regard

to the chain length of the alcohol.32 Transesteri®cation

1538 J Sci Food Agric 79:1535±1549 (1999)

FD Gunstone

of soy phospholipids at the sn-1 position occurs with

methyl esters of C10±C18 acids (Mucor miehei) to

furnish phospholipids with capric (8%), lauric (14%),

myristic (16%), or linolenic acid (5% raised to 15%).33

5 FATTY ACIDSThere has been an increasing awareness in recent years

of the nutritional importance of certain fatty acids such

as g-linolenic acid, arachidonic acid, and docosahex-

aenoic acid. These occur naturally but often only at

low levels along with other acids. There is therefore a

demand for preparations with higher levels of these

acids and, at least for research purposes, for prepara-

tions of virtually pure material. While such material

can be produced by conventional physical and

chemical methods of separation these are not always

appropriate if the material is subsequently to be used

for nutritional studies. This problem has been

successfully overcome by exploiting the speci®city of

those enzymes which discriminate against acids having

double bonds at position 4 as in docosahexaenoic acid,

position 5 as in arachidonic acid, or position 6 as in

g-linolenic acid) The concentration or isolation of such

acids is discussed in the following sections.

(i) g-Linolenic acidg-Linolenic acid (GLA) is an important acid used as a

dietary supplement and available from three major

vegetable sources: the seed oils of evening primrose

(8±10%), borage (20±25%), and blackcurrant (15±

17%). These levels of GLA can be raised by enzymic

enhancement and evening primrose and borage oils

with about twice the original GLA levels are commer-

cially available. Laboratory procedures have been

described by Hills et al34 using lipase from rape

seedlings or Mucor miehei and by Foglia and Sonnet35

using Geotrichum candidum lipase. Procedures are

illustrated in the recent study of Shimada et al.36

Borage oil (22% GLA) is hydrolysed (Pseudomonasspecies, 35°C, 24h) and the acids are selectively

esteri®ed with lauric alcohol (Rhizopus delemar, 30°C,

20h). The latter enzyme discriminates against GLA

which therefore concentrates in the unesteri®ed acids.

These ®nally contain 70% GLA (74% recovery). If the

esteri®cation is repeated, 94% GLA is obtained with

68% recovery. Concentration of GLA by selective

deacylation of the oils is also possible but is less

ef®cient. Shimada et al37 showed that selective

hydrolysis of borage oil (22% GLA) gave glycerol

esters with 46% of this acid (C rugosa, 35°C, 15h)

which could hardly be raised above this value. After

removal of free acids, a second hydrolysis raised the

concentration to 54% with 76% recovery. On a larger

scale 7kg of borage oil gave 1.5kg of upgraded oil

(56% GLA) after removal of free acid by molecular

distillation. Starting with an already upgraded borage

oil (10kg, 45% GLA) high quality g-linolenic acid

(2.1kg, 98% pure, 49% recovery) was obtained by

hydrolysis and two separate partial esteri®cations.38

(ii) Arachidonic acidThe single cell oil from Mortierella alpina is now com-

mercially available as a useful triacylglycerol source of

arachidonic acid containing no n-3 acids. Partial

hydrolysis of this oil with an enzyme which discrimi-

nates against arachidonic acid, leads to enrichment of

this acid in the glycerol ester fraction. When oil con-

taining 25% of arachidonic acid is partially hydrolysed

(Candida antarctica, 35°C, 16h, 52% hydrolysis) the

glycerol ester fraction is mainly triacylglycerols with

50% arachidonic acid (95% recovery). If the process is

repeated the level of arachidonic acid rises to 54%

(88% recovery) and 60% (75% recovery).39 (Shimada

and Sugihara et al39). In a later report Shimada et al40

raised the level of arachidonic acid to 75% using the

two-step process already reported for g-linolenic

acid.36 The oil (25% arachidonic acid) is hydrolysed

to free acids (Pseudomonas sp, 40°C, 40h). Subse-

quently these are esteri®ed selectively with lauric

alcohol (Candida rugosa, 30°C, 16h, 55% esteri®ca-

tion) to give residual free acids with 51% arachidonic

acid (92% recovery). This is raised to 63% (78%

recovery) by urea fractionation and to 75% (71%

recovery) by a second esteri®cation.

(iii) Docosahexaenoic acidSeveral procedures for enzymic enhancement of n-3

PUFA, and especially DHA, have been reported. In

general, ®sh oils are subject to selective hydrolysis or

alcoholysis (ethanol or lauryl alcohol) or free acids are

partially esteri®ed (ethanol or butanol). Several

enzymes have given useful results based on the fact

that they discriminate against DHA which therefore

concentrates in the residual substrate.

Selective hydrolysis of seal blubber oil and menha-

den oil (Candida cylindracea, 30±50°C, 46h) gives

glycerol esters enriched in n-3 acids (Table 3).41

McNeill et al42 and Moore et al43 used lipases from

Candida rugosa and Geotrichum candidum to raise levels

of EPA and DHA from 20±30% in several ®sh oils to

35±45% in partial glycerol esters. These can be con-

verted to triacylglycerols by hydrolysis followed by

reaction with glycerol, both reactions being catalysed

by Rhizopus miehei.Shimada et al44 raised DHA levels of tuna ethyl

esters by selective alcoholysis with lauric alcohol from

23 to 50±52% (Rhizomucor miehei or Rhizopus dele-mar). Higher levels were obtained by starting with

already enriched material (45 raised to 74% and 60

raised to 93%). EPA levels are reduced slightly during

Table 3. Enzymic enrichment of polyunsaturated fatty acids in fish oils

Seal blubber oil Menhaden oil

EPA DPA DHA EPA DPA DHA

Original oil 6.4 4.7 7.6 13.2 2.4 10.1

Residual esters 9.8 8.6 24.0 18.5 3.6 17.3

Enrichment 1.5 1.8 3.2 1.4 1.5 1.7

J Sci Food Agric 79:1535±1549 (1999) 1539

Enzymes as biocatalysts in the modi®cation of lipids

this reaction. This result is expected on the basis of

reaction rates relative to those of ethyl oleate (=100):

EPA 13 and 24, DHA 15 and 11 for Rhizomucordelemar and Rhizopus miehei respectively.

Attempts to raise levels of EPA and DHA have been

described by Haraldsson et al45,46 and Breivik et al.47

These are summarised in Table 4. The same authors

have described ways of separating EPA and DHA by

kinetic resolution using Rhizomucor miehei lipase.

(iv) Erucic acidSonnet et al48 concentrated erucic acid from a high-

erucic rape seed oil (HERO) by complete hydrolysis of

the oil (Pseudomonas cepacia) followed by selective

esteri®cation with butanol (Geotrichum candidum).

This latter enzyme discriminates against erucic acid

and its level was raised from 48% in HERO to �85%

in the unesteri®ed acids.

6 Alkyl estersAlkyl esters based on primary (ethyl, butyl) or

secondary (isopropyl, iso-octyl) alcohols can be made

by esteri®cation of free acids or by transesteri®cation

of glycerol (or other) esters using enzymic catalysts.

Such a reaction is frequently one step in a series of

processes for the concentration of individual acids (eg

Refs 44, 46). Methods have been described for

obtaining esters on a kilogram scale and as a step in

upgrading low-quality material such as acid oil or

soapstock.

Ethyl stearate, for example, has been prepared in

high yield from technical stearic acid in 2kg batches

using Lypozyme.49 Esters were also made in high yield

on the 2kg scale by esteri®cation of oleic acid with

butanol and by transesteri®cation of rape seed oil with

iso-octanol using the lipases from Candida rugosa,Chromobacterium viscosum, Pseudomonas ¯uorescens, or

Rhizopus miehei.50

Acid oils containing a mixture of free acids (40±

80%) and glycerol esters (20±50%) can be hydrolysed

(Candida cylindracea lipase, 35°C, 48h) to acids, or the

acid/ester mixture can be converted to alkyl esters (4:0

to 18:0 alcohols) with Mucor miehei lipase.23 Soapstock

(water 45%, acids 10%, glycerol esters 12%, and

phospholipids 9%) can also be upgraded by chemical

hydrolysis followed by reaction with alcohol (methyl,

ethyl, butyl, isobutyl, tert-amyl) or by transesteri®ca-

tion using Lipozyme or Candida antarctica lipase at

42°C.51

Lipase-catalysed transesteri®cation has been recom-

mended for the preparation of methyl or ethyl (Mucormiehei) and isopropyl or isobutyl (Candida antarctica)

esters from tallow or vegetable oils for use as

biodiesel.52 Similar reactions can be carried out using

diesel fuel as solvent to give mixtures of diesel and

biodiesel.53

Deodoriser distillate54 is a valuable by-product of

the re®ning process which contains free acids along

with the more valuable tocopherols and sterols. To

recover the two latter the acids must be removed and

this can be achieved by distillation after enzyme-

catalysed conversion to methyl esters (Randozyme,

50°C) in yields up to 96%.

7 MONO- AND DI-ACYLGLYCEROLSMonoacylglycerols, sometimes with diacylglycerols,

are important food-grade emulsi®ers used as such or

after further chemical modi®cation. Estimates of

annual world production vary between 0.25 and 0.50

million tonnes. They are produced mainly by alkali-

catalysed glycerolysis of oils and fats (220±260°C,

30min). The product is a mixture of monoacylglycer-

ols (45±55%), diacylglycerols (38±45%), and triacyl-

glycerols (8±12%). The monoacylglycerol content can

be increased to over 90% through short-path distilla-

tion. The high temperatures involved in this process

make it less suitable for unsaturated compounds. The

monoacylglycerol is an equilibrium mixture of the 1-

(90%) and 2- (10%) isomers.55,56

Enzymic routes to monoacylglycerols and diacylgly-

cerols hace been studied extensively. They include

interaction of glycerol with acid, alkyl ester, or

triacylglycerol or reaction of triacylglycerol with water

(partial hydrolysis), alkyl alcohol (partial alcoholysis)

or glycerol (glycerolysis). These procedures have been

reviewed.57 Under appropriate reaction conditions it is

possible to obtain products with high levels of

monoacylglycerols without the short-path distillation

required in the existing commercial method.

McNeill et al58±61 showed that high levels of

monoacylglycerols were obtained by glycerolysis of

triacylglycerols when the reaction was carried out at

appropriately low temperatures. Monoacylglycerols

have higher melting points than other glycerol esters

and may crystallise from the reaction mixture thereby

displacing the equilibrium in favour of more mono-

acylglycerol. The reaction temperature has to be a little

below the melting point of the monoacylglycerol. This

is higher for saturated acids (52±84°C for C10 to C22)

than for unsaturated acids (oleic 35°C, linoleic

12°C).55 For example, glycerolysis of beef tallow (rich

in palmitic and stearic acids) at 48±50°C gives a

product with 30% monoacylglycerols but at 38±46°Cthis is raised to 70%. Neither solvent nor emulsi®er is

required. Pseudomonas ¯uorescens lipase is an effective

catalyst but others have also been used. Better yields

Table 4. Enrichment of EPA and DHA in fish oils

EPA, DHA

(%) Lipasea Hours Product

EPA,

DHA (%)

Tuna oil 6 and 23 Rm 24 gl esters 6 and 49

Tuna acids 6 and 23 Rm 8 acids 3 and 74

Fish oil 15 and 9 Ps 24 gl esters (50)b

a Rm=Rhizomucor miehei Ps=Pseudomonas sp.b The EPA�DHA level rises to 85% when the glycerol esters are converted to

ethyl esters (Candida antarctica) and submited to urea fractionation.

1540 J Sci Food Agric 79:1535±1549 (1999)

FD Gunstone

are obtained in a two-stage process in which reaction is

started at ca42°C for 16h and continued at 5°C for up

to four days. Under these conditions tallow, palm oil

and palm stearin were converted to monoacylglycerols

in yields of 90±95%. Similar reactions have been

reported for palm oil,62 camphor tree seed oil (a lauric

oil,63), cocoa butter,63 marine oils,64 olive oil,65,66

soybean oil67 and triolein.68

Yamane et al69 adapted the procedure to give high

yields (90%) of diacylglycerols in a two-phase glycer-

olysis of hydrogenated beef tallow effected at 60°C(2h), 55°C (4h), and then 48°C for three days.

An alternative route to mono- and diacylglycerols

involves direct reaction of glycerol with free acid or

some other source of acyl group. Macrae et al70

recommended the use of potato lipid acyl hydrolase to

promote reaction (50±70°C, 6h) between glycerol and

free acid (10:0, 12:0, 14:0, 18:1, 18:2, or 18:3) to give

monoacylglycerols in yields above 95%. Berger et al71

describe the acylation of glycerol on silica (organic

solvent, room temperature, 2±5h, in the presence of

Chromobacterium viscosum, Rhizopus delemar or Rhizo-mucor miehei lipases) with a range of acids (5:0±18:0

and 18:1). 1,3-Diacylglycerols were produced in

80±90% yield and isolated with purity >98%. The

same research group72 optimised procedures to get

high yields of monoacylglycerols (60±85%) using

excess of glycerol (1:5) and of diacylglycerols

(50±75%) using excess of acid (2:1) with the lipases

from Penicillium roquefortii or Rhizopus niveus at room

temperature. Akoh et al73 prepared monoacylglycerols

from glycerol and oleic acid (or ethyl oleate) or from

glycerol and eicosapentaenoic acid in yields of 90±98%

(Penicillium sp, molecular sieve, 25±37°C, one to four

days). Dicaprin was obtained from glycerol and ethyl

caprate in a solvent-free system in yields up to 88%

(Rhizopus arrhizus, 15±50°C).74 Edmundo et al75 pre-

pared mono-olein (71% conversion, 100% pure after

passage through a silica column) from glycerol on

silica, oleic acid, and Mucor miehei lipase in 2-methyl-

2-butanol/hexane (10:90) at 40°C.

8 TRIACYLGLYCEROLSThere is a growing demand for compounds which are

described as structured lipids. These are triacylglycer-

ols containing speci®c fatty acids in designated

positions. Laboratory processes for synthesising in-

dividual triacylglycerols are well developed76 but these

are not appropriate to produce these materials in the

volume required for nutritional purposes even in a

research exercise. Nor is it usually necessary to

produce compounds which are entirely a single species

± concentrates will usually suf®ce. Using an appro-

priate lipase as catalyst it is possible to prepare such

materials by acylation of glycerol or by deacylation/

reacylation of existing triacylglycerol mixtures. The

most important of these contain a medium-chain acid

and/or a long-chain polyunsaturated fatty acid.

Enzymic acylation of glycerol with free acid or alkyl

ester proceeds readily but the product may be mainly a

monoacylglycerol/diacylglycerol mixture rather than

triacylglycerol. This is caused by the dif®culty in

acylating the sn-2 hydroxyl group by direct acylation or

(more likely) through acyl migration.

Glycerol → 1(3)-MAG → 1,3-DAG → TAG

;1,2-DAG → TAG

(all these processes must be considered reversible)

An example is given in Table 5 for reaction between

glycerol and n-3 PUFA.77

This dif®culty does not exist when the starting point

for triacylglycerol synthesis is 2-monoacylglycerol

though care must be taken that reaction is con®ned

to acylation and that both acyl-migration and trans-

esteri®cation are kept to a minimum. This is illustrated

in the synthesis of caprucin from 2-monoerucin and

caprylic acid (Geotrichum candidum, 50°C, 75%

yield78) and in the two-step synthesis of triacylglycer-

ols described by Soumanou et al79,80 (Section 8(i)).

(i) Medium chain triacylglycerolsMedium-(M) and long-chain (L) acyl groups are

metabolised differently and since each type shows

valuable nutritional properties there is an interest in

preparing triacylglycerols with both types of acyl chain,

particularly those represented as MLM where the long

acyl chain occupies the glycerol sn-2 position. The

preparation of such compounds has been studied

intensively by Akoh and his research group. Using

Rhizopus miehei and Candida antarctica, they examined

the acidolysis and interesteri®cation of triolein, tri-

linolein, and tristearin with acids (C4, C6, and C10),

ethyl esters (C8), and triacylglycerols (C6 and C8) in

organic solvent and showed that in all cases the

resulting triacylglycerols were mainly mixtures of the

type L2M and LM2. With R miehei exchange was

con®ned almost entirely to the sn-1 and 3 positions but

with C antarctica reaction also occurred at the sn-2

position.81±87 Lee and Akoh88 studied the solvent-free

acidolysis of groundnut oil and caprylic acid in a batch

reactor on a 500g scale. The highest incorporation of

caprylic acid (c 30%) was achieved with a 1:2 molar

ratio at 50°C for 72h with Rhizomucor miehei. When

tricaprylin is submitted to acidolysis with n-3 PUFA

(Candida antarctica, 55°C, hexane) followed by short-

path distillation, structured lipids are obtained con-

taining caprylic acid (47%), eicosapentaenoic acid

(23%), and docosahexaenoic acid (22%) present in all

three of the glycerol positions.89 Capric acid and

Table 5. Composition of reaction product from glycerol andn-3 PUFA

MAG DAG TAG FFA (%)

Pseudomonas sp 24 41 18 17

Mucor miehei 34 41 13 12

J Sci Food Agric 79:1535±1549 (1999) 1541

Enzymes as biocatalysts in the modi®cation of lipids

eicosapentaenoic acid were both incorporated into

borage seed oil (already containing g-linolenic acid) in

hexane solution using the enzymes from Rhizomucormiehei and Candida antarctica as biocatalysts. Signi®-

cant proportions of both acids are incorporated but

they are found in the sn-2 position only when Candidaantarctica lipase is employed. (Table 6, Ref 90).

Kwon et al91 examined the enzymic reaction of

glycerol and capric acid (iso-octane, 25°C, up to 25h).

Non-selective lipases gave less satisfactory results than

1,3-selective lipases but not all of this latter type gave

the same results. Better results were obtained with

Rhizomucor miehei, Pseudomonas aeruginosa or Chromo-bacterium viscosum lipase) than with Aspergillus niger,Rhizopus javanicus or Rhizopus delemar lipase. The

major products were triacylglycerols but monoacylgly-

cerols and diacylglycerols were also present in the

reaction mixture. A range of PUFA were converted to

glycerol esters with lipases from Chromobacteriumviscosum (26±38% TAG) and Candida cylindracea(18±33% TAG) and more successfully with those

from Rhizomucor miehei and Candida cylindracea at

40±60°C (80±90%).92 Kosugi and Azuma93 reported

that immobilised lipase from Candida antarcticapromoted acylation of glycerol with arachidonic acid,

eicosapentaenoic acid, or docosahexaenoic acid or

their ethyl esters at 60°C. After 24h the product was

mainly TAG (80±90%) and was easily puri®ed by

passage through an alumina column. See also Refs 42

and 43.

Acylation of glycerol on silica gel with capric acid in

iso-octane at 40±60°C has been examined with a range

of lipases. Tricaprin was the major product with some

lipases (Pseudomonas aeruginosa, Rhizomucor miehei,and Chromobacterium viscosum) and dicaprin with

other lipases (Candida rugosa and Rhizopus delemar).94

With Rhizomucor miehei (60°, 5h) Ghosh et al95 raised

the levels of caprylic and capric acids in coconut oil or

coconut olein by interesteri®cation with the methyl

esters of these short-chain acids from ca12 to 20±25%.

Gioielli et al96 have described the modi®cation of

babassu oil (a lauric oil) by acidolysis with palmitic

acid (raised from 10 to 22%, Rhizomucor miehei, 65°C,

4h).

A group working in Stuttgart79,80,97 have shown that

MLM triacylglycerols are best made in a two-step

process. Triacylglycerols (triolein, trilinolein, or

groundnut oil) are ®rst subject to ethanolysis (Rhizo-pus delemar, 40°C, 30h, MeOBut-hexane solution) to

give a 2-monoacylglycerol. This is puri®ed by crystal-

lisation from hexane (72% yield of recovered product)

and then acylated (Rhizomucor miehei, caprylic acid,

hexane, molecular sieve, 38°C) to give a ®nal product

containing caprylic acid (94%) in the sn-1 and 3

positions and unsaturated C18 acid (98%) in the sn-2

position.

Xu et al98,99 have described studies to optimise the

acidolysis reaction (®sh oil and capric acid or medium-

chain triglycerides and sun¯ower acids) in a one-

kilogram reactor with emphasis on the minimisation of

acyl migration.

(ii) Triacylglycerols enriched in EPA and/or DHAStructured lipids containing eicosapentaenoic acid

and/or docosahexaenoic acid can be made in several

ways: (i) by reaction of glycerol with polyunsaturated

fatty acids or their ethyl esters, (ii) by enrichment of

acids already present in ®sh oils by appropriate

deacylation and reacylation procedures, and (iii) by

reaction of vegetable or other oils with PUFA con-

centrates as free acids, ethyl esters or glycerol esters.

Acids/esters enriched in eicosapentaenoic acid and/or

docosahexaenoic acid are often obtained from ®sh oils

by urea fractionation of the acids or alkyl esters.

Glycerol and a concentrate of polyunsaturated fatty

acids (24% EPA and 53% DHA) reacted in organic

solvent under the in¯uence of Chromobacterium visco-sum lipase to give a mixture of monoacylglycerols

(14%, with 27% EPA and 50% DHA), diacylglycerols

(43%, with 25% EPA and 50% DHA), and triacylgly-

cerols (37%, with 21% EPA and 50% DHA).100

Cerdan et al101 used the lipase from Candida antarcticato convert glycerol and a PUFA concentrate (26%

EPA and 48% DHA) from cod liver oil into a product

which was 85% triacylglycerol with 26% EPA and

45% DHA (60°C, 96h).

Several papers describe the concentration of eico-

sapentaenoic acid and docosahexaenoic acid in ®sh

oils by partial hydrolysis with a lipase that discrimi-

nates against DHA (and EPA). These acids concen-

trate in the residual glycerol esters which are generally

mixtures of di- and triacylglycerols that can be acylated

to triacylglycerols. Reference has already been made to

work by a number of researchers41±47 on the concen-

tration of docosahexaenoic acid (Section 5). Tanaka etal102 converted tuna oil (25% DHA) by partial

hydrolysis (Candida antarctica) followed by acylation

with PUFA from tuna oil (Chromobacterium viscosum,

50°) to triacylglycerols with 39±46% of DHA. Maehr

et al103 used lipase from Pseudomonas sp at ambient

temperature to raise the level of n-3 PUFA from 29±

34% to 50% with 23±50% recovery or to over 70%

with 14±22% recovery.

Yamane et al104,105 used the lipase from Mucormiehei or Candida cylindracea to raise the levels of EPA

and DHA in cod liver oil (9 and 13%) to 22 and 34%

when the reaction was conducted at ÿ10°C for 20h

followed by ÿ20°C for 40h.

Sridhar et al106 incorporated EPA (9%) and DHA

Table 6. Distribution of selected acids in borage oil afteracidolysisa

10:0 18:2 g-18:3 20:5

Borage ± 39 (47) 20 (20) ±

Modi®ed Rm 26 (±) 21 (49) 17 (22) 10 (±)

Modi®ed Ca 16 (8) 27 (50) 17 (15) 9 (5)

a Levels of 10:0 and 20:5 in the treated borage oil

(percentage in the sn-2 position).

1542 J Sci Food Agric 79:1535±1549 (1999)

FD Gunstone

(8%) into groundnut oil using PUFA with 35 and 20%

of these two acids and Mucor miehei lipase for 4±6h.

Akoh and his colleagues85,107,108 incorporated EPA

and/or DHA into several vegetable oils (canola,

groundnut, soybean, hydrogenated soybean and

high-oleic sun¯ower), and trilinolein, tricaprylin,

tricaprin, and trilaurin with Mucor miehei and Candidaantarctica lipases at 55°C for 24h. Lee et al85 showed

that these enzymes incorporate 35±42% of EPA into

short- and medium-chain triacylglycerols but they

behave differently (Table 7). With Mucor miehei EPA

was con®ned to the sn-1(3) position but with Candidaantarctica it occurred in all three glycerol positions.

(iii) Margarine and shorteningMargarines and shortenings made with partially

hydrogenated fats contain acids with trans unsatura-

tion which are out of favour on nutritional grounds.

The melting behaviour required of a spread can be

achieved, at some small cost in terms of increased

saturated acids, by interesteri®cation of suitable blends

of hard stock and vegetable oils and this change is

usually effected using an alkaline catalyst.109,110

Similar changes can be achieved with appropriate

lipases and several procedures have been described.

For example, a product with melting behaviour

appropriate for use as a margarine or shortening was

made by enzymic interesteri®cation of a high-melting

palm stearin (mp 58°C) with a vegetable oil (sun-

¯ower, soybean, or rice bran) in 40:60 ratio (Mucormiehei, 60°C, 5h). The product contained polyunsa-

turated fatty acids and was virtually free of transacids.111 When lard and high-oleic sun¯ower (60:40)

were interesteri®ed (Candida antarctica, 55°C, 24h)

the product was superior to that obtained by blending

alone and had solid content resembling that of a soft

margarine at a range of temperatures.112 From the

examination of a ternary mixture of canola oil,

hydrogenated canola oil, and palm stearin Cho and

deMan113 concluded that the interesteri®cation pro-

duct crystallised in the desirable b' form and could be

used to produce margarine with trans levels lower than

conventional products made from hydrogenated oils

even though trans acids were still present.

Foglia et al114 have reported changes in melting

behaviour resulting from enzymic interesteri®cation of

tallow, high-oleic sun¯ower oil, or tallow-sun¯ower

mixtures (Mucor miehei, Rhizopus delemar, or Geo-trichum candidum at 50±70°C). They conclude that

glyceride composition can be in¯uenced by choice of

enzyme under controlled conditions in contrast to

chemical reaction products that are wholly random in

nature.

Graille et al115 have studied the interesteri®cation of

palm oil and palm stearin with several other oils using a

1,3-regiospeci®c lipasae (Mucor miehei, 60°C). They

obtained products which, depending on the conditions

employed, can be used for ®rm or soft margarines.

(iv) Cocoa butter equivalentsEnzymic interesteri®cation of oils and fats to produce

cocoa butter equivalents has been reviewed by

Quinlan and Moore116 and by Rosendaal and

Macrae109 and is covered by patents on these materials

to Unilever and to Fuji Oil Co. The production of

useful products has been described by Bloomer et al117

(palm mid fraction and ethyl stearate), Chang etal118(hydrogenated cottonseed oil and olive oil), and

by Liu et al119 (palm oil and tristearin).

9 WAX ESTERS, SUGAR ESTERS, AND ESTERSOF POLYHYDRIC ALCOHOLS OTHER THANGLYCEROLLipases from Rhizopus miehei or Candida antarcticahave been used to promote acylation of medium-chain

(8:0) and long-chain (18:0) alcohols with the acyl

group provided as free acid, methyl ester, or triacyl-

glycerol. Reaction occurs at ca 60°C and high yields

(95±100%) of wax esters are obtained in 30±

120min.120±123 Krmelj et al124 examined the lipase-

catalysed synthesis of oleyl oleate in pressurised and

supercooled solvents using Rhizomucor miehei lipase.

In butane 87% conversion was obtained (20°C, 20

bar, 5h) while carbon dioxide gave 86% conversion in

one hour (50°C, 100 bar).

Hayes and Kleiman125 described the Lypozyme-

catalysed reaction between diols and either lesquerella

oil or lesquerolic acid to give mono- and diesters which

have potential application as cosmetics and as lubri-

cants. The secondary hydroxyl group in lesquerolic

acid is not involved in the process.

Sugar estersIt is possible to prepare lipid-like molecules by

replacing glycerol with mono- or disaccharides. The

acylated sugars have interesting nutritional properties

and can also be used as surfactants.126 The best known

is olestra, a sucrose derivative with six to eight

esteri®ed groups obtained from sucrose and common

vegetable oils (soybean, corn, cottonseed, sun¯ower,

rapeseed). It is prepared by reaction of high-grade

sucrose with high-grade methyl esters in the presence

of sodium or potassium soaps to improve solubility

and an alkali carbonate as catalyst. It can be used as a

frying oil and can replace fats in products such as ice

cream, margarine, cheese and baked goods. The

material is non-toxic, non-carcinogenic, and is so

poorly absorbed as to have zero calori®c value.

Permission has been granted for limited use in the

Table 7. Incorporation of EPA into short- and medium-chaintriacylglycerolsa

Mucor miehei Candida antarctica

Tricaprylin 34 (±) 38 (35)

Tricaprin 32 (±) 42 (39)

Trilaurin 36 (±) 41 (36)

a Values of EPA incorporated into the glycerol ester (EPA in

the sn-2 position).

J Sci Food Agric 79:1535±1549 (1999) 1543

Enzymes as biocatalysts in the modi®cation of lipids

USA and on balance it has been well received but it is

not yet licensed for use elsewhere.

Partly acylated sugars with a balance of hydrophobic

and hydrophilic groups show interesting surfactant

properties. Attempts to produce such molecules by

standard chemical syntheses are likely to be tedious

and instead, attempts have been made to exploit the

speci®city of lipase. Mutua et al127 described the

acylation of methyl glucoside and related alky glyco-

sides with methyl oleate (Candida sp, 55°C, 24±48h,

benzene-pyridine) with 60±100% incorporation of the

acyl group. The same research group acylated glucose

penta-acetate with methyl oleate or high-oleic sun-

¯ower oil as the acylating agent in the presence of

lipases from Candida antarctica and Candida rugosa.128

In a more controlled procedure Sarney et al129

achieved the regioselective synthesis of sucrose

monoesters (6'O-acyl- and 6-O-acyl) by lipase-cata-

lysed acylation (Candida antarctica and Rhizomucormiehei, 48h, 75°C, range of organic solvents) of

appropriate sucrose acetals in which secondary hydro-

xyl groups are blocked leaving the primary hydroxyl

groups free for acylation.

The Stuttgart group130 has shown that drawbacks

like low solubility, low reaction rate, and low

productivity associated with the enzymic biotransfor-

mations of polar sugars in dilute organic solvents can

be easily overcome even when carrying out these

reactions in a system with solid substrates and

products. The correct choice and concentration of

the organic solvent serving as adjuvant is most

important to obtain high yields of sugar fatty acid

esters. Typical reaction conditions leading to good

results (up to 100% acylation of primary hydroxyl

group) require acetone, Candida antarctica, 12:0±18:0

fatty acids, 48h. A clear advantage of this reaction

system is the use of readily available starting materials

and no activated acyl donors or modi®ed sugars in

order to achieve high conversions in acceptable

reaction times.

Other estersAcylated derivatives of propylene glycol (1,2-propane

diol) are mixtures of mono-(mainly 1-) and di-esters

and are used as food-grade surfactants. With propy-

lene glycol, Pseudomonas lipase, and organic solvent

(hexane, toluene) at 30°C the product was mainly the

mono-ester obtained in yields of 29±37% (with acids),

65±86% (with triacylglycerols), and 80±90% (with

anhydrides) depending on the nature of the C12±C18

acyl group.131 The same group have prepared propy-

lene glycol esters containing EPA and DHA.132

Trimethylolpropane (TMP) esters, used as bio-

degradable lubricants, are normally made through

high-temperature acylation. They can, however, be

obtained through alcoholysis of rapeseed methyl esters

with TMP using lipase from Candida rugosa.133

The glycerol ether 1-O-hexadecylglycerol can be

acylated by C14±C18 fatty acids in the presence of

Candida cylindracea or Pseudomonas sp lipases (di-

chloromethane, three days at 33°C or for a shorter

time at 55°C) to give a 75% yield of a 7:1 mixture of

the 3- and 2-acyl ethers.134

10 AmidesLipases from Rhizopus miehei and Candida antarcticahave been used to promote acylation of amino

compounds to give amides using triacylglycerols or

methyl esters as the source of the acyl group. Examples

include the N-acylation of lysine,135 butylamine and

other amines,136 and b-alanine.137

11 CHEMICAL REACTIONS OF THE ALKYL CHAINGardner138 reviewed the role of lipoxygenase as a

versatile biocatalyst and Gargouri and Legoy139

illustrated its use in the preparation of (�)-coriolic

acid (13S-OH 9c11t-18:2) from trilinolein or from

sun¯ower oil used as a rich source of linoleic acid. The

triacylglycerols were ®rst hydrolysed by lipase from

Pseudomonas sp to the free acids which were then

oxidised by lipoxygenase to the 13-hydroperoxy acid

and reduced chemically (NaBH4-EtOH) to the hydro-

xy acid (purity >95%, yield 17±78% depending on

concentration of triacylglycerol). Reaction occurs at

pH 9 and 28°C in 150±200min in a two-phase system.

Hydrolysis occurs at the interface and oxidation in the

aqueous phase.

ÐCH=CHCH2CH=CHÐ →

CH(OOH)CH=CHCH=CHÐ →

CH(OH)CH=CHCH=CHÐ

By reaction with hydrogen peroxide in toluene in the

presence of Candida antarctica lipase triacylglycerols or

methyl esters are converted to peroxy acids which then

effect epoxidation of unsaturated centres.140 With

rape, sun¯ower, and linseed oils epoxidation is about

90% complete. Frykman and Isbell141 applied a

similar procedure to meadowfoam fatty acids, rich in

D5 acids (30% hydrogen peroxide in toluene or

dichloromethane and Candida antarctica lipase) and

subsequently converted the epoxy acids to hydroxy

lactones and estolides by chemical processes.

Bacillus lentus NRRL B-14864 converts 12-hydro-

xystearic acid (from ricinoleic acid) to a C12 tetra-

hydrofuran acid, via chain-shortening (b-oxidation) to

6-hydroxydodecanoic acid. This intermediate under-

goes cyclisation at the 2t or 3-hydroxy stage during the

next oxidation cycle.142

In the presence of appropriate enzyme sources, oleic

acid and other appropriate acids are converted to a

range of oxidation products which are not easily

produced with chemical reagents. Oleic acid is

1544 J Sci Food Agric 79:1535±1549 (1999)

FD Gunstone

converted to 15-, 16-, and 17-hydroxyoleic acids (5±

11%) with Bacillus pumilis in three to ®ve days at

32°C143 and to 7R,10R-dihydroxy 8t-18:1 with

bacterial strain PR3.144 With Pseudomonas aeruginosaPR3 the yield of dihydroxy acid was 80%145 and this

compound can be reduced with hydrazine to 7R,10R-

dihydroxystearic acid. Linoleic acid did not react in

this system but ricinoleic acid gave 7,10,12-trihydroxy

8t-18:1 (35% yield) by an analogous process.

Flavobacterium sp DS5 hydratase converts oleic acid

to a mixture of 10-hydroxy and 10-oxo-stearic acid.

Other D 9c acids (9c-14:1, 9c-16:1, linoleic, and a- and

g-linolenic acids) react similarly to give appropriate

10-hydroxy and oxo acids but there is no reaction with

elaidic (9t-18:1), erucic (13c-22:1), arachidonic (all-cis5,8,11,14±20:4), or petroselinic (6c-18:1) acids, all of

which lack the D 9 cis double bond.146

ÐCH=CHÐ → ÐCHOHCH2Ð → ÐCOCH2Ð

The microbial culture Clavibacter sp ALA2 converts

linoleic acid to 12,13,17-trihydroxy 9c-18:1 as major

product (25%) accompanied by minor products (each

3±4%) which have been identi®ed as the tetrahydro-

furan derivatives: 12-hydroxy- and 7,12-dihydroxy-

13,16-epoxy 9c-18:1.147,148

12 REVIEWSSeveral aspects of this topic have been reviewed pre-

viously. Malcata149 described the use of immobilised

enzymes in the modi®cation of oils and fats by

hydrolysis, ester synthesis, and interesteri®cation.

Immobilisation procedures, reactor con®guration,

and process considerations are also discussed. Born-

scheuer57 has described the lipase-catalysed synthesis

of monoacylglycerols, and the ability of lipases to

convert hydroxy acids to lactones and estolides is

considered by Hayes.150 Gandhi151 has produced a

useful review on lipase applications covering both

hydrolysis and ester synthesis. Other reviews devoted

to lipases are by Quinlan and Moore116 and by

Rosendaal and Macrae.109

Uhlig152 in his book ªIndustrial Enzymes and theirApplicationsº includes a section on lipases, including

animal lipases, microbial lipases, and phospholipases.

The most recent review of this topic is a chapter on

enzymic processes by McNeill153 who is also an author

of several signi®cant papers cited in this review. After a

discussion of the enzymes themselves, his review is

focused on potential products (triacylglycerols, di-

acylglycerols, monoacylglycerols, fatty acids, phos-

pholipids, sugar esters, and waxes).

In his review of the role of lipoxygenase as a versatile

biocatalyst Gardner138 includes details of the chemical

and enzymic reactions of hydroperoxides, the latter

with allene oxide synthase and hydroperoxide lyase.

Over 100 compounds have been identi®ed from

linoleic acid and related compounds are obtained

from other PUFA.

13 CONCLUSIONInterest in the use of lipases and other enzymes to

promote reactions, some of which are not easily

effected by non-enzymic processes, has grown con-

siderably in the last 10 years in both academic and

industrial laboratories. This is re¯ected in a rapidly

growing number of publications. At the present time

interest is focussed on the use of these systems to

concentrate and isolate polyunsaturated fatty acids

required for dietary studies in animals and in humans

and on the production of structured lipids. The latter

are triacylglycerols (and phospholipids) with speci®c

fatty acids or fatty acid classes esteri®ed with glycerol

in speci®c positions generally on the basis of their

nutrirional properties. One can expect further devel-

opments in the next ten years and the arrival of more

products in the market place.

REFERENCES1 Gunstone FD, Movements towards tailor-made fats. Prog Lipid

Res 37:277±305 (1998).

2 Hilditch TP, The Industrial Chemistry of Fats and Waxes, Balliere,

Tindall and Cox, London, pp 235±237 (1927).

3 Holmquist M, Insights into the molecular basis for fatty acyl

speci®cities of lipases from Geotrichum candidum and Candida

rugosa. Chem Phys Lipids 93:57±65 (1998).

4 Lang DA and Dijkstra BW, Structural investigations of the

regio- and enantioselectivity of lipases. Chem Phys Lipids

93:115±122 (1998).

5 Klein RR, King G, Moreau RA and Haas MJ, Altered acyl chain

length speci®city of Rhizopus delemar lipase through mutagen-

esis and molecular modeling. Lipids 32:123±130 (1997).

6 Carriere F, Withers-Martinez C, van Tilbeurgh H, Roussel A,

Cambillau C and Verger R, Structure-function relationships

of pancreatic lipases. Fett/Lipid 100:96±102 (1998).

7 Murukami M, Kawasaki Y, Kawanari M and Okai H, Trans-

esteri®cation of oil by fatty acid-modi®ed lipase. J Am Oil

Chem Soc 70:571±574 (1993).

8 Mogi K-I and Nakajama M, Selection of surfactant-modi®ed

lipases for transesteri®cation of triglyceride and fatty acid. J

Am Oil Chem Soc 73:1505±1512 (1996).

9 Sonnet PE and Gazzillo JA, Evaluation of lipase selectivity for

hydrolysis. J Am Oil Chem Soc 68:11±15 (1991).

10 Sonnet PE, Foglia TA and Baillargeon MW, Fatty acid

selectivity: the selectivity of lipases of Geotrichum candidum.

J Am Oil Chem Soc 70:1043±1045 (1993).

11 Sonnet PE, McNeill GP and Jun W, Lipase of Geotrichum

candidum immobilised on silica gel. J Am Oil Chem Soc

71:1421±1423 (1994).

12 Gulomova K, Ziomek E, Schrag JD, Davranov K and Cygler M,

Puri®cation and characterisation of a Penicillium sp lipase

which discriminates against diglycerides. Lipids 31:379±384

(1996).

13 Osterberg E, Blomstrom A-C and Holmberg K, Lipase-

catalysed transesteri®cation of unsaturated lipids in a micro

emulsion. J Am Oil Chem Soc 66:1330±1333 (1989).

14 Lee I and Hammond EG, Oat (Avena sativa) caryiopses as a

natural lipase bioreactor. J Am Oil Chem Soc 67:761±765

(1990).

15 Piazza GJ, Bilyk A, Brower DP and Haas MJ, The positional

and fatty acid selectivity of oat seed lipase in aqueous

emulsions. J Am Oil Chem Soc 69:978±981 (1992).

16 Parmar S and Hammond EG, Hydrolysis of fats and oils with

moist oats caryopses. J Am Oil Chem Soc 71:881±886 (1994).

17 Hou CT and Johnston TM, Screening of lipase activities with

J Sci Food Agric 79:1535±1549 (1999) 1545

Enzymes as biocatalysts in the modi®cation of lipids

cultures from the Agricultural Research Service culture

collection. J Am Oil Chem Soc 69:1088±1097 (1992).

18 Hou CT, Characterization of new yeast lipases. J Am Oil Chem

Soc 74:1391±1394 (1997).

19 Castillo E, Dossat V, Marty A, Condoret JS and Combes D,

The role of silica gel in lipase-catalysed esteri®cation reactions

of high-polar substrates. J Am Oil Chem Soc 74:77±85 (1997).

20 Hass MJ, Cichowicz DJ, Jun W and Scott K, The enzymatic

hydrolysis of triglyceride-phospholipid mixtures in an organic

solvent. J Am Oil Chem Soc 72:519±525 (1995).

21 Fu X, Zhu X, Gao K and Duan J, Oil and fat hydrolysis with

lipase from Aspergillus sp. J Am Oil Chem Soc 72:527±531

(1995).

22 Kosugi Y and Tomizuka N, Continuous lipolysis reactor with a

loop connecting an immobilised lipase column and an oil-

water separator. J Am Oil Chem Soc 72:1329±1332 (1995).

23 Ghosh S and Bhattacharyya DK, Utilization of acid oils in

making valuable fatty products by microbial lipase technol-

ogy. J Am Oil Chem Soc 72:1541±1544 (1995).

24 Virto MD, Lascaray JM, Solozabal R and de Renobales M,

Enzymic hydrolysis of animal fats in organic solvents at

temperatures below their melting point. J Am Oil Chem Soc

68:324±327 (1991).

25 Na A, Eriksson C, Eriksson S-G, Osterberg E and Holmberg K,

Synthesis of phosphatidylcholine with (n-3) fatty acids by

phospholipase A2 in microemulsion. J Am Oil Chem Soc

67:766±770 (1990).

26 Haas MJ, Scott K, Jun W and Janssen G, Enzymatic phos-

phatidylcholine hydrolysis in organic solvents: an examina-

tion of selected commercially available lipases. J Am Oil Chem

Soc 71:483±490 (1994).

27 Hara F, Nakashima T and Fukuda H, Comparative study of

commercially available lipases in hydrolysis reaction of phos-

phatidylcholine. J Am Oil Chem Soc 74:1129±1132 (1997).

28 Ono M, Hosokawa M, Inoue Y and Takahashi K, Water

activity-adjusted enzymatic partial hydrolysis of phospho-

lipids to concentrate polyunsaturated fatty acids. J Am Oil

Chem Soc 74:1415±1417 (1997).

29 Svensson I, Adlercreutz P and Mattiasson B, Lipase-catalyzed

transesteri®cation of phosphatidylcholine at controlled water

activity. J Am Oil Chem Soc 69:986±991 (1992).

30 Mustranta A, Suortti T and Poutanen K, Transesteri®cation of

phospholipids in different reaction conditions. J Am Oil Chem

Soc 71:1415±1419 (1994).

31 Sarney DB, Fregapane G and Vulfson EN, Lipase-catalyzed

synthesis of lysophospholipids in a continuous bioreactor. J

Am Oil Chem Soc 71:93±96 (1994).

32 Ghosh M and Bhattacharyya DK, Enzymatic alcoholysis

reaction of soy phospholipids. J Am Oil Chem Soc 74:597±

599 (1997).

33 Ghosh M and Bhattacharyya DK, Soy lecithin-monoester

interchange reaction by microbial lipase. J Am Oil Chem Soc

74:761±763 (1997).

34 Hills MJ, Klewitt I and Mukherjee KD, Enzymatic fractiona-

tion of fatty acids: enrichment of g-linolenic acid and

docosahexaenoic acid by selective esteri®cation catalyzed by

lipases. J Am Oil Chem Soc 67:561±564 (1990).

35 Foglia TA and Sonnet PE, Fatty acid selectivity of lipases: g-linolenic acid from borage oil. J Am Oil Chem Soc 72:417±420

(1995).

36 Shimada Y, Sugihara A, Shibahiraki M, Fujita H, Nakano H,

Nagao T, Terai H and Tominaga Y, Puri®cation of g-linolenic acid from borage oil by a two-step enzymatic

method. J Am Oil Chem Soc 74:1465±1470 (1997).

37 Shimada Y, Fukushima N, Fujita H, Honda Y, Sugihara A and

Tominaga Y, Selective hydrolysis of borage oil with Candida

rugosa lipase: two factors affecting the reaction. J Am Oil

Chem Soc 75:1581±1586 (1998).

38 Shimada Y, Sakai N, Sugihara A, Fujita H, Honda Y and

Tominaga Y, Large-scale puri®cation of g-linolenic acid by

selective esteri®cation using Rhizopus delemar lipase. J Am Oil

Chem Soc 75:1539±1543 (1998).

39 Shimada Y, Sugihara A, Maruyama K, Nagao T, Nakayama S,

Nakano H and Tominaga Y, Enrichment of arachidonic acid:

selective hydrolysis of a single cell oil from Mortierella with

Candida cylindracea lipase. J Am Oil Chem Soc 72:1323±1327

(1995).

40 Shimada Y, Sugihara A, Minamigawa Y, Higashiyama K,

Akimoto K, Fujikawa S, Komenmushi S and Tominaga Y,

Enzymatic enrichment of arachidonic acid from Mortierella

single cell oil. J Am Oil Chem Soc 75:1213±1217 (1998).

41 Wanasundara UN and Shahidi F, Lipase-assisted concentration

of n-3 polyunsaturated fatty acids in acylglycerols from

marine oils. J Am Oil Chem Soc 75:945±951 (1998).

42 McNeill GP, Ackman RG and Moore SR, Lipase-catalyzed

enrichment of long-chain polyunsaturated fatty acids. J Am

Oil Chem Soc 73:1403±1407 (1996).

43 Moore SR and NcNeill GP, Production of triglycerides

enriched in long-chain n-3 polyunsaturated fatty acids from

®sh oil. J Am Oil Chem Soc 73:1409±1414 (1996).

44 Shimada Y, Maruyama K, Sugihara A, Baba T, Komemushi S,

Moriyama S and Tominaga Y, Puri®cation of ethyl docosa-

hexaenoate by selective alcoholysis of fatty acid ethyl esters

with immobilized Rhizopus miehei lipase. J Am Oil Chem Soc

75:1565±1571 (1998).

45 Haraldsson GG, Kristinsson B, Sigurdardottir R, Gudmunds-

son GG and Breivik H, The preparation of concentrates of

eicosapentaenoic acid and docosahexaenoic acid by lipase-

catalyzed transesteri®cation of ®sh oil with ethanol. J Am Oil

Chem Soc 74:1419±1424 (1997).

46 Haraldsson GG and Kristinsson B, Separation of eicosapentae-

noic acid and docosahexaenoic acid by kinetic resolution

using lipase. J Am Oil Chem Soc 75:1551±1556 (1998).

47 Breivik H, Haraldsson and Kristinsson B, Preparation of highly

puri®ed concentrates of eicosapentaenoic acid and docosa-

hexaenoic acid. J Am Oil Chem Soc 74:1425±1429 (1997).

48 Sonnet PE, Foglia TA and Feairheller SH, Fatty acid selectivity

of lipases: erucic acid from rapeseed oil. J Am Oil Chem Soc

70:387±391 (1993).

49 Bloomer S, Adlercreutz P and Mattiasson B, Kilogram-scale

ester synthesis of acyl donor and use in lipase-catalyzed

interesteri®cations. J Am Oil Chem Soc 69:966±973 (1992).

50 Linko Y-Y, Limsa M, Huhtala A and Rantanen O, Lipase

biocatalysis in the production of esters. J Am Oil Chem Soc

72:1293±1299 (1995).

51 Haas MJ and Scott KM, Combined nonenzymatic-enzymatic

method for the synthesis of simple alkyl fatty acid esters from

soapstock. J Am Oil Chem Soc 73:1393±1401 (1996).

52 Nelson LA, Foglia TA and Marmer WN, Lipase-catalyzed

production of biodiesel. J Am Oil Chem Soc 73:1191±1195

(1996).

53 Haas MJ and Scott KM, Diesel fuel as a solvent for the lipase-

catalyzed alcoholysis of triglycerides and phosphatidylcho-

line. J Am Oil Chem Soc 73:1497±1504 (1996).

54 Ramamurthi S, Bhirud PR and McCurdy AR, Enzymatic

methylation of canola oil deodorizer distillate. J Am Oil Chem

Soc 68:970±975 (1991).

55 Krog N, Food emulsi®ers, in Lipid Technologies and Applications

ed by Gunstone FD, Marcel Dekker, New York. pp 421±534

(1997).

56 Hassenheuttl GL, Synthesis and commercial preparation of

surfactants in the food industry, in Lipid Synthesis and

Manufacture, ed by Gunstone FD, Shef®eld Academic Press,

Shef®eld, UK. pp 371±400 (1999).

57 Bornscheuer U, Lipase-catalysed synthesis of monoglycerides.

Fat Sci Technol 97:241±249 (1995).

58 McNeill GP, Shimizu S and Yamane T, Solid phase enzymatic

glycerolysis of beef tallow resulting in a high yield of mono-

glyceride. J Am Oil Chem Soc 67:779±783 (1990).

59 McNeill GP, Shimizu S and Yamane T, High-yield enzymatic

glycerolysis of fats and oils. J Am Oil Chem Soc 68:1±5 (1991).

1546 J Sci Food Agric 79:1535±1549 (1999)

FD Gunstone

60 McNeill GP, and Yamane T, Further improvements in the yield

of monoglycerides during enzymatic glycerolysis of fats and

oils. J Am Oil Chem Soc 68:6±10 (1991).

61 McNeill GP, Borowitz E and Berger RG, Selective distribution

of saturated fatty acids into the monoglyceride fraction during

enzymatic glycerolysis. J Am Oil Chem Soc 69:1098±1103

(1992).

62 Holmberg K and Osterberg E, Enzymatic preparation of

monoglycerides in microemulsion. J Am Oil Chem Soc

65:1544±1548 (1988).

63 Thude S, Shukun L, Said MB and Bornscheuer UT, Lipase-

catalyzed synthesis of monoacylglycerides by glycerolysis of

camphor tree seed oil and cocoa-butter. J Am Oil Chem Soc

99:246±250 (1997).

64 Myrnes B, Barstad H, Olsen RL and Elvevoll EO, Solvent-free

enzymatic glycerolysis of marine oils. J Am Oil Chem Soc

72:1339±1344 (1995).

65 Rosu R, Uozaki Y, Iwasaki Y and Yamane T, Repeated use of

immobilised lipase for monoacylglycerol production by solid-

phaase glycerolysis of olive oil. J Am Oil Chem Soc 74:445±450

(1997).

66 Coteron A, Martinez M and Aracil J, Reactions of olive oil and

glycerol over immobilized lipases. J Am Oil Chem Soc 75:657±

660 (1998).

67 Jackson MA and King JW, Lipase-catalyzed glycerolysis of

soybean oil in supercritical carbon dioxide. J Am Oil Chem Soc

74:103±106 (1997).

68 Arcos JA and Otero C, Enzyme, medium and reaction

engineering to design a low-cost, selective production method

for mono- and dioleylglycerols. J Am Oil Chem Soc 73:673±

682 (1996).

69 Yamane T, Tang ST, Kawahara K and Koizumi Y, High-yield

diacylglycerol formation by solid-phase enzymatic glyceroly-

sis of hydrogenated beef tallow. J Am Oil Chem Soc 71:339±

342 (1994).

70 Macrae AR, Visicchio JE and Lanot A, Application of potato

lipid acyl hydrolase for the synthesis of monoacylglycerols. J

Am Oil Chem Soc 75:1489±1494 (1998).

71 Berger M, Laumen K and Schneider MP, Enzymatic esteri®ca-

tion of glycerol I. Lipase-catalyzed synthesis of regioiso-

merically pure 1,3-sn-diacylglycerols II Lipase-catalyzed

synthesis of regioisomerically pure 1(3)rac monoacylglycer-

ols. J Am Oil Chem Soc 69:955±960 and: 961±965 (1992).

72 Waldinger C and Schneider M, Enzymatic esteri®cation of

glycerol III. Lipase-catalysed synthesis of regioisomerically

pure 1,3-diacylglycerols and 1(3)-rac-monoacylglycerols de-

rived from unsaturated acids. J Am Oil Chem Soc 73:1513±

1519 (1996).

73 Akoh CC, Cooper C and Nwosu CV, Lipase G-catalyzed

synthesis of monoglycerides in organic solvent and lipid

analysis by HPLC. J Am Oil Chem Soc 69:257±260 (1992).

74 Fureby AM, Adlercreutz P and Mattiasson B, Glyceride

synthesis in a solvent-free system. J Am Oil Chem Soc

73:1489±1495 (1996).

75 Edmundo C, Valerie D, Didier C and Alain M, Ef®cient lipase-

catalyzed production of tailor-made emulsi®ers using solvent

engineering coupled to extractive processing. J Am Oil Chem

Soc 75:309±313 (1998).

76 Sonnet PE, Synthesis of triacylglycerols, in Lipid Synthesis and

Manufacture, ed by Gunstone FD, Shef®eld Academic Press,

Shef®eld, UK. pp 162±184 (1999).

77 Li Z-Y and Ward OP, Lipase-catalyzed esteri®cation of glycerol

and n-3 polyunsaturated fatty acid concentrate in organic

solvent. J Am Oil Chem Soc 70:745±748 (1993).

78 McNeill GP and Sonnet PE, Low-calorie triglyceride synthesis

by lipase-catalyzed esteri®cation of monoglycerides. J Am Oil

Chem Soc 72:1301±1307 (1995).

79 Soumanou MM, Bornscheuer UT and Schmid RD, Two-step

enzymatic reaction for the synthesis of pure structured

triglycerides. J Am Oil Chem Soc 75:703±710 (1998).

80 Soumanou MM, Bornscheuer UT, Schmid U and Schmid RD,

Synthesis of structured triglycerides by lipase catalysis. Fett/

Lipid 100:156±160 (1998).

81 Akoh CC and Yee LN, Enzymatic synthesis of position-speci®c

low-calorie structured lipids. J Am Oil Chem Soc 74:1409±

1413 (1997).

82 Fomuso LB and Akoh CC, Enzymatic modi®cation of triolein:

incorporation of caproic and butyric acids to produce

reduced-calorie structured lipids. J Am Oil Chem Soc

74:269±272 (1997).

83 Fomuso LB and Akoh CC, Structured lipids: lipase-catalyzed

interesteri®cation of tricaproin and trilinolein. J Am Oil Chem

Soc 75:405±410 (1998).

84 Huang K-H and Akoh CC, Enzymatic synthesis of structured

lipids: transesteri®cation of triolein and caprylic acid ester. J

Am Oil Chem Soc 73:245±250 (1996).

85 Lee K-T and Akoh CC, Immobilised lipase-catalyzed produc-

tion of structured lipids with eicosapentaenoic acid at speci®c

positions. J Am Oil Chem Soc 73:611±615 (1996).

86 Lee K-T and Akoh CC, Effects of selected substrate forms on

the synthesis of structured lipids by two immobilized lipases. J

Am Oil Chem Soc 74:579±584 (1997).

87 Sheih C-J, Akoh CC and Koehler PE, Four-factor response

surface optimization of the enzymatic modi®cation of triolein

to structured lipids. J Am Oil Chem Soc 72:619±623 (1995).

88 Lee K-T and Akoh CC, Solvent-free enzymatic synthesis of

structured lipids from peanut oil and caprylic acid in a stirred

tank batch reactor. J Am Oil Chem Soc 75:1533±1537 (1998).

89 Lee K-T and Akoh CC, Characterization of enzymatically

synthesized structured lipids containing eicosapentaenoic,

docosahexaenoic, and caprylic acids. J Am Oil Chem Soc

75:495±499 (1998).

90 Akoh CC and Mousata CO, Lipase-catalyzed modi®cation of

borage oil: incorporation of capric and eicosapentaenoic acids

to form structured lipids. J Am Oil Chem Soc 75:697±701

(1998).

91 Kwon DY, Song HN and Yoon SH, Esteri®cation patterns of

lipases for synthesizing tricaproylglycerols in organic solvent.

J Am Oil Chem Soc 74:1287±1290 (1997).

92 Osada K, Takahashi K and Hatano M, Polyunsaturated fatty

glyceride syntheses by microbial lipases. J Am Oil Chem Soc

67:921±922 (1990).

93 Kosugi Y and Azuma N, Synthesis of triacylglycerol from

polyunsaturated fatty acid by immobilised lipase. J Am Oil

Chem Soc 71:1397±1403 (1994).

94 Kwon DY, Song HN and Yoon SH, Synthesis of medium-chain

glycerides by lipase in organic solvent. J Am Oil Chem Soc

73:1521±1525 (1996).

95 Ghosh S and Bhattacharyya DK, Medium-chain fatty acid-rich

glycerides by chemical and lipase catalyzed polyester-monoe-

ster interchange reaction. J Am Oil Chem Soc 74:593±595

(1997).

96 Gioielli LA, Pitombo RNM, Baruffaidi R, Oliveira MN and

Augusto MS, Acidolysis of babassu fat catalyzed by im-

mobilized lipase. J Am Oil Chem Soc 71:579±582 (1994).

97 Soumanou MM, Bornscheuer UT, Menge U and Schmid RD,

Synthesis of structured triglycerides from peanut oil with

immobilized lipase. J Am Oil Chem Soc 74:427±433 (1997).

98 Xu X, Skands ARJ, Adler-Nissen J and Hoy C-E, Production of

speci®c lipids by enzymatic interesteri®cation: optimization of

the reaction by response surface design. Fett/Lipid 100:463±

471 (1998).

99 Xu X, Balchen S, Hoy C-E and Adler-Nissen J, Pilot batch

production of speci®c-structured lipids by lipase-catalyzed

interesteri®cation: preliminary study on incorporation and

acyl migration. J Am Oil Chem Soc 75:301±308 (1998).

100 He Y and Shahidi F, Enzymatic esteri®cation of o-3 fatty acid

concentrates from seal blubber oil with glycerol. J Am Oil

Chem Soc 74:1133±1136 (1997).

101 Cerdan LE, Medina RA, Gimenez AG, Gonzalez MJI and

Grima EM, Synthesis of polyunsaturated fatty acid-enriched

J Sci Food Agric 79:1535±1549 (1999) 1547

Enzymes as biocatalysts in the modi®cation of lipids

triglycerides by lipase-catalyzed esteri®cation. J Am Oil Chem

Soc 75:1329±1337 (1998).

102 Tanaka Y, Hirano J and Funada T, Synthesis of docosahex-

aenoic acid-rich triglyceride with immobilised Chromobacter-

ium viscosum lipase. J Am Oil Chem Soc 71:331±334 (1994).

103 Maehr H, Zenchoff G and Coffen DL, Enzymic enhancement

of n-3 fatty acid content in ®sh oils. J Am Oil Chem Soc

71:463±467 (1994).

104 Yamane T, Suzuki T, Sahashi Y, Vikersveen L and Hoshino T,

Production of n-3 polyunsaturated fatty acid-enriched ®sh oil

by lipase-catalyzed acidolysis without solvent. J Am Oil Chem

Soc 69:1104±1107 (1992).

105 Yamane T, Suzuki T and Hoshino T, Increasing n-3

polyunsaturated fatty acid content of ®sh oil by temperature

control of lipase-catalyzed alcoholysis. J Am Oil Chem Soc

70:1285±1287 (1993).

106 Sridhar R and Lakshminarayana G, Incorporation of eicosa-

pentaenoic and docosahexaenoic acids into groundnut oil by

lipase-catalyzed ester interchange. J Am Oil Chem Soc

69:1041±1042 (1992).

107 Akoh CC, Jennings BH and Lillard DA, Enzymatic modi®ca-

tion of trilinolein: incorporation of n-3 polyunsaturated fatty

acids. J Am Oil Chem Soc 72:1317±1321 (1995).

108 Huang K-H and Akoh CC, Lipase-catalyzed incorporation of

n-3 polyunsaturated fatty acids into vegetable oils. J Am Oil

Chem Soc 71:1277±1280 (1994).

109 Rosendaal A and Macrae AR, Interesteri®cation of oils and fats,

in Lipid Technologies and Applications ed by Gunstone FD and

Padley FB, Dekker pp 223±263 (1997).

110 Allan D, Fat modi®cation as a tool for product development.

Part 2: Interesteri®cation and biomodi®cation. Lipid Technol-

ogy 10:53±57 (1998).

111 Ghosh S and Bhattacharyya DK, Utilization of high-melting

palm stearin in lipase-catalyzed interesteri®cation with liquid

oils. J Am Oil Chem Soc 74:589±592 (1997).

112 Seriburi V and Akoh CC, Enzymatic interesteri®cation of lard

and high-oleic sun¯ower oil with Candida antarctica lipase to

produce plastic fats. J Am Oil Chem Soc 75:1339±1345

(1998).

113 Cho F and deMan JM, Physical properties and composition of

low-trans canola/palm blends modi®ed by continuous enzy-

matic interesteri®cation. Elaies 6:39±48 (1994).

114 Foglia TA, Petruso K and Feairheller SH, Enzymatic inter-

esteri®cation of tallow-sun¯ower oil mixtures. J Am Oil Chem

Soc 70:281±285 (1993).

115 Graille J, Pina M, Montet D and Muderhwa JM, Making value-

added products from palm oil by 1±3 regioselectivity

enzymatic interesteri®cation. Elais 4:1±10 (1991).

116 Quinlan P and Moore S, Modi®cation of triglycerides by lipases

process technology and its application to the production of

nutritionally improved fats. INFORM 4:580±585 (1993).

117 Bloomer S, Adlercreutz P and Mattiasson B, Triglyceride

interesteri®cation by lipases. I Cocoa butter equivalents from

a fraction of palm oil. J Am Oil Chem Soc 67:519±524 (1990).

118 Chang M-K, Abraham G and John VT, Production of cocoa

butter-like fat from interesteri®cation of vegetable oils. J Am

Oil Chem Soc 67:832±834 (1990).

119 Liu K-J, Cheng H-M, Chang R-C and Shaw J-F, Synthesis of

cocoa butter equivalent by lipase-catalyzed interesteri®cation

in supercritical carbon dioxide. J Am Oil Chem Soc 74:1477±

1482 (1997).

120 Trani M, Ergan F and Andre G, Lipase-catalyzed production of

wax esters. J Am Oil Chem Soc 68:20±22 (1991).

121 Multzsch R, Lokotsch W, Steffen B, Lang S, Metzger JO,

Schafer HJ, Warwel S and Wagner F, Enzymatic production

and physicochemical characterisation of uncommon wax

esters and monoglycerides. J Am Oil Chem Soc 71:721±725

(1994).

122 Goma-Doncescu N and Legoy MD, An original transesteri®-

cation route for fatty acid ester production from vegetable oils

in a solvent-free system. J Am Oil Chem Soc 74:1137±1143

(1997).

123 MuÈhlhaus R, Jupke A and Mehrwald A, Process development of

fatty acid esteri®cation catalyzed enzymatically through

process simulation. Fett/Lipid, 99:392±399 (1997).

124 Krmelj V, Habulin M, Knez Z and Bauman D, Lipase-catalyzed

synthesis of oleyl oleate in pressurized and supercritical

solvents. Fett/Lipid 101:34±38 (1999).

125 Hayes DG and Kleiman R, Lipase-catalyzed synthesis of

lesquerolic acid wax and diol esters and their properties. J

Am Oil Chem Soc 73:1385±1392 (1996).

126 Sakidja and Swanson BG, Alkyl and acyl sugars, in Lipid

Synthesis and Manufacture, ed by Gunstone FD, Shef®eld

Academic Press, Shef®eld, UK. pp 347±370 (1999).

127 Mutua LN and Akoh CC, Synthesis of alkyl glycoside fatty acid

esters in non-aqueous media by Candida sp lipase. J Am Oil

Chem Soc 70:43±46 (1993).

128 Akoh CC, Enzymatic synthesis of acetylated glucose fatty acid

esters in organic solvent. J Am Oil Chem Soc 71:319±323

(1994).

129 Sarney DB, Barnard MJ, MacManus DA and Vulfson EN,

Application of lipases to the regioselective synthesis of sucrose

fatty acid monoesters. J Am Oil Chem Soc 73:1481±1487

(1996).

130 Cao L, Bornscheuer UT and Schmid RD, Lipase-catalyzed

solid phase synthesis of sugar esters. Fett/Lipid 98:332±335

(1996).

131 Shaw J-F and Lo S, Production of propylene glycol fatty acid

monoesters by lipase-catalysed reactions in organic solvents.

J Am Oil Chem Soc 71:715±719 (1994).

132 Liu K-J and Shaw J-F, Synthesis of propylene glycol monoesters

of docosahexaenoic acid and eicosapentaenoic acid by lipase-

catalyzed esteri®cation in organic solvents. J Am Oil Chem Soc

72:1271±1274 (1995).

133 Uosukainen E, Linko Y-Y, Lamsa M, Tervakangas T and Linko

P, Transesteri®cation of trimethylolpropane and rapeseed oil

methyl ester to environmentally acceptable lubricants. J Am

Oil Chem Soc 75:1557±1563 (1998).

134 Bertello LE, Salto ML and de Lederkremer RM, Lipase-

catalyzed monoesteri®cation of 1-O-hexadecylglycerol in

organic solvents. Lipids 32:907±911 (1997).

135 Montet D, Servat F, Pina M, Graille J, Galzy P, Arnoud A,

Ledon H and Marcou L, Enzymatic synthesis of N-e-acyllysines. J Am Oil Chem Soc 67:771±774 (1990).

136 Bistline RG, Bilyk A and Feairheller SH, Lipase-catalyzed

formation of fatty amides. J Am Oil Chem Soc 68:95±98

(1991).

137 Izumi T, Yagimuma Y and Haga M, Enzymatic syntheses of

N-lauroyl-b-alanine homologs in organic media. J Am Oil

Chem Soc 74:875±878 (1997).

138 Gardner HW, Lipoxygenase as a versatile biocatalyst. J Am Oil

Chem Soc 73:1347±1357 (1996).

139 Gargouri M and Legoy MD, Chemoenzymatic production of

(�)-coriolic acid from trilinolein: coupled synthesis and

extraction. J Am Oil Chem Soc 74:641±645 (1997).

140 Rusch gen Klaas M and Warwel S, Chemoenzymatic epoxida-

tion of unsaturated fatty acid esters and plant oils. J Am Oil

Chem Soc 73:1453±1457 (1996).

141 Frykman HB and Isbell TA, Synthesis of 6-hydroxy d-lactones

and 5,6-dihydroxy eicosanoic/docosanoic acids from mea-

dowfoam fatty acids via a lipase-mediated self-epoxidation. J

Am Oil Chem Soc 74:719±722 (1997).

142 Huang J-K, Dowd PF, Keudell KC, Klopfenstein WE, Wen L,

Bagby MO, Lanser AC and Norton RA, Biotransformation

of saturated monohydroxyl fatty acids to 2-tetrahydrofuranyl

acetic acid derivatives: mechanism of formations and

biological activity of 5-n-hexyl-tetrahydrofuran-2-acetic acid.

J Am Oil Chem Soc 73:1465±1469 (1996).

143 Lanser AC, Plattner RD and Bagby MO, Production of 15-,

16-, and 17-hydroxy-9-octadecenoic acid by bioconversion of

1548 J Sci Food Agric 79:1535±1549 (1999)

FD Gunstone

oleic acid with Bacillus pumilus. J Am Oil Chem Soc 69:363±

366 (1992).

144 Knothe G, Bagby MO, Peterson RE and Hou CT, 7,10-

Dihydroxy-8(E)-octadecenoic acid: sterochemistry and a

novel derivative, 7,10-dihydroxy-octadecanoic acid. J Am

Oil Chem Soc 69:367±371 (1992).

145 Kuo TM, Manthey LK and Hou CT, Fatty acid bioconversions

by Pseudomonas aeruginosa PR3. J Am Oil Chem Soc 75:875±

879 (1998).

146 Hou CT, Production of hydroxy fatty acids from unsaturated

fatty acids by Flavobacterium sp DS5 hydratase, a C-10

positional- and cis unsaturation-speci®c enzyme. J Am Oil

Chem Soc 72:1265±1270 (1995).

147 Hou CT, A novel compound, 12,13,17-trihydroxy-9(Z)-

octadecenoic acid from linoleic acid by a new microbial

isolate Clavibacter sp ALA2. J Am Oil Chem Soc 73:1359±

1362 (1996).

148 Hou CT, Gardner H and Brown W, Production of polyhydroxy

fatty acids from linoleic acid by Clavibacter sp ALA2. J Am Oil

Chem Soc 75:1483±1487 (1998).

149 Malcata FX, Reyes HE, Garcia HS, Hill CG and Amundsen

CH, Immobilized lipase reactors for modi®cation of fats and

oils±a review. J Am Oil Chem Soc 67:890±910 (1990).

150 Hayes DG, The catalytic activity of lipases toward hydroxy fatty

acids±a review. J Am Oil Chem Soc 73:543±549 (1996).

151 Gandhi NN, Applications of lipase. J Am Oil Chem Soc 74:621±

634 (1997).

152 Uhlig H, Industrial Enzymes and their Applications, Wiley, New

York. pp 179±192 (1998).

153 McNeill GP, Enzymic processes, in Lipid Synthesis and

Manufacture ed by Gunstone FD, Shef®eld Academic Press,

Shef®eld, UK. pp 288±320 (1999).

J Sci Food Agric 79:1535±1549 (1999) 1549

Enzymes as biocatalysts in the modi®cation of lipids