ARTICLE
Auteur(s) : Céline Tiran1,2, Jérôme
Lecomte1, Eric
Dubreucq2, Pierre
Villeneuve1
1CIRAD, UMR 1208 IATE, Montpellier, F-34398
France
2Montpellier SupAgro, UMR 1208 IATE, Equipe
Biotechnologie microbienne et enzymatique des lipides et des
agropolymères, 2 Place Viala, F-34060 Montpellier cedex 1,
France
The worldwide potential demand for replacing petroleum-derived
raw materials with renewable ones for the production of polymeric
materials is quite significant from the social and environmental
points of view [1]. Beside polysaccharides and sugars, plant oils
are the most important renewable raw materials of the chemical
industry because of their ready availability [2].
Among the major applications of epoxidized vegetable oils is
their use as plasticizer for polyvinyl chloride (PVC) and other
plastic materials [3]. Plasticizers are substances that improve
flexibility, workability or distensibility of plastics, hence
rendering them suitable for diverse applications. Epoxy fatty acids
can also be used as PVC-stabilisers because of their ability to
slow down degradation by scavenging the free HCl released during
PVC decomposition when exposed to heat and light [4]. In addition,
epoxidized derivatives of fatty acids can be used as reactive
diluents for paints, as intermediates for polyurethane-polyol
production, as corrosion protecting agents and as additives to
lubricating oils. They also represent valuable raw materials for
the production of glues and other surface coatings [5].
Epoxidized oils are currently produced by epoxidation of
unsaturated plant oils, such as soybean or linseed oil. Although
several methods are available to epoxidize the double bonds of
unsaturated fatty acids, the only method applied on industrial
scale is the Prileshajev epoxidation reaction. In this reaction,
short chain peroxy acids such as peracetic acid are generated from
the corresponding acid and hydrogen peroxide in the presence of a
strong mineral acid. Then, these peroxy acids react with
unsaturated fatty acid C=C double bonds to obtain epoxidized fatty
acids. Peroxy acids are prepared either in a separate step or in
situ. Due to the potential danger of handling peroxy acids, the in
situ method is generally preferred for large-scale epoxidation of
unsaturated triglycerides [6].
However, this chemical method for epoxidation has several
disadvantages. First of all, there are considerable side reactions
via oxirane ring opening, leading to diols, hydroxyesters,
estolides and other dimers, which are believed to be catalyzed by
the presence of a strong mineral acid [7]. As a result, the
selectivity of this process never exceeds 80% [6]. Furthermore, the
presence of a strong acid in an oxidative environment causes
equipment corrosion problems. Finally, this acid must be recycled
or neutralized before discharge into the environment. Using enzymes
is an alternative solution that allows an environmentally benign
and more selective epoxidation reaction.
Main enzymes involved in epoxidation
Several enzyme types are directly or indirectly involved in fatty
acid epoxidation: cytochrome P450 monooxygenases, diiron-center
oxygenases, lipoxygenases, peroxygenases, perhydrolases and
lipases. They belong to the oxydoreductase (EC 1.x.x.x) or
hydrolase (EC 3.x.x.x) enzyme classes.
Oxydoreductases
Cytochrome P450 monooxygenases (EC 1.14.x.y) are one of the
largest and oldest enzyme superfamilies with several thousand
sequences reported up to now [5, 8]. Cytochromes P450 contain a
heme-thiolate prosthetic group. They are widely distributed in
animals, plants, fungi and bacteria and catalyze a vast range of
monooxygenation reactions in catabolic and anabolic pathways [9,
10]. P450s incorporate a single atom of molecular oxygen into a
substrate with the concomitant reduction of the other atom to
water. With respect to fatty acids, they can act both as
hydroxylases and as epoxidases of unsaturated fatty acids. The
reducing equivalents are delivered by the nicotinamide cofactor
NAD(P)H. As this cofactor is far too expensive for industrial
applications, one major challenge for all attempts to construct a
bioreactor with isolated P450 enzymes is to engineer an artificial
electron supply system, generally consisting in a coupling with a
deshydrogenase-catalyzed NAD(P) reduction [11]. CYP102A1, also
called P450 BM-3, is one of the most intensely studied P450
monooxygenases. This 119 kDa enzyme, originally cloned from
Bacillus megaterium, is a self-sufficient P450 in which an FAD- and
FMN-containing reductase and the P450-domain are naturally fused on
a single peptide chain [12]. The experimental setup for its
application in organic synthesis is a lot easier compared to other
P450s which require one or two additional electron transport
proteins for activity. Indeed, in the presence of NADPH and
O2 P450 BM-3 can catalyze the oxygenation of long chain
fatty acids without the aid of any other protein. Moreover, all
self-sufficient P450 monooxygenases characterized to date exhibit
rather high turnover numbers (> 1,000 s-1) with
their preferred substrates [5].
Diiron-center oxygenases are mainly present in plants and
bacteria. They are often referred to as “diiron-oxo” proteins
because most proteins containing a binuclear diiron cluster react
with dioxygen as part of their functional processes [13]. These
proteins catalyze diverse reactions including hydroxylation,
epoxidation, and desaturation according to the following catalytic
mechanism: reductive oxygen is activated by the diiron-center, a
hydrogen atom from a CH-bond is then abstracted and the oxygen atom
is incorporated into the substrate via a radical rebound mechanism
[5, 14]. Diiron-center oxygenases and desaturases share high
sequence homology. From a biotechnological point of view,
diiron-oxo oxygenases present the same disadvantages that P450
enzymes as both enzyme classes use electrons originating from the
costly cofactor NAD(P)H for reductive oxygen activation and require
an electron transport chain delivering these reduction equivalents.
That is why, despite their occurrence in most plant species
producing oils enriched in epoxy acids, attemps to exploit diiron
cluster-containing fatty acid oxygenases in biotechnology are still
scarce.
Lipoxygenases (EC 1.13.11.12; LOXs) constitute a large
gene family of non-heme iron containing fatty acid dioxygenases,
which are ubiquitous in plants and animals. They catalyze the
regio- and stereospecific incorporation of dioxygen into
polyunsaturated fatty acids (linoleic acid, α-linolenic acid or
arachidonic acid) to generate optically active (S)-dienic
hydroperoxides [15-17]. Plant lipoxygenases are classified
according to their regioselectivity using linoleic acid as
substrate. Linoleic acid is oxygenated either at the Δ9 carbon atom
(9-LOX) or at the Δ13 carbon atom (13-LOX) of the hydrocarbon
backbone of the fatty acid leading to the formation of two groups
of compounds: the (9S)-hydroperoxy- and the (13S)-hydroperoxy
derivatives (figure
1). For example, soybean LOX isoenzyme-1 is classified as a
13-LOX whereas potato tuber LOX is classified as a 9-LOX [5]. Both
the cavity within the active site and the orientation of the
substrate are important determinants for the regiospecificity of
plant lipoxygenases [18]. The lipid hydroperoxides formed by
lipoxygenases are highly reactive and are immediately used by plant
cells for the biosynthesis of a wide range of compounds including
epoxides. The metabolism of oxidized polyunsaturated fatty acids
via the lipoxygenase-catalyzed step and the subsequent reactions
are collectively called the LOX pathway [16, 19, 20]. Concerning
biotechnological applications, the use of isolated lipoxygenases in
organic solvents and their immobilization have up to now been
poorly investigated [21].
Peroxygenases (EC 1.14.x.y) are P450-related enzymes of
the LOX pathway that use the hydroperoxides formed by lipoxygenases
as substrate. These enzymes are membrane-bound proteins containing
heme b as prosthetic group [22] and are ubiquitous in plants. The
first reactions described for peroxygenases are co-oxidative
reactions such as epoxidation, hydroxylation, and sulfoxidation,
where an oxygen atom from the hydroperoxide is directly transferred
to the substrate. When using hydroperoxides deriving from
unsaturated fatty acids, this oxygen transfer can also occur via an
intramolecular mechanism [23]. Peroxygenases do not require
cofactors such as NAD(P)H or FAD. In the presence of an organic
hydroperoxide, oleic acid is converted into the corresponding
9,10-epoxide by peroxygenase activities isolated from soybean,
broad bean and oat [23-25]. Piazza et al. [26, 27] have developed a
method for the rapid isolation and immobilization of peroxygenases
on membranes, and conducted epoxidation reactions in organic
solvents.
Hydrolases
Hydrolases such as lipases (EC 3.1.1.3) and carboxyl
esterases (EC 3.1.1.1) have been shown to produce peroxy acids
from hydrogen peroxide and fatty acids by perhydrolysis reaction.
These peroxy acids subsequently epoxidize unsaturated fatty acids
via an uncatalyzed reaction. In this case, large-scale applications
seem to be possible in the near future. The next chapter of this
review focuses on lipase-catalyzed perhydrolysis.
Lipase-catalyzed perhydrolysis
Chemo-enzymatic epoxidation of oils and fats
In 1990, an immobilized form of lipase B from Candida antarctica
(Novozyme 435) was shown to catalyze the conversion of saturated
fatty acids into peroxy fatty acids in the presence of hydrogen
peroxide [28]. Indeed, perhydrolysis, where hydrogen peroxide acts
as the nucleophile instead of water in the deacylation step, might
be a side activity of many serine hydrolases. However, the
perhydrolase activity of lipases and esterases is generally much
lower than their esterase activity and some of them, such as
subtilisin, do not exhibit perhydrolase activity [29]. To the
contrary, some hydrolases display a higher perhydrolysis activity
than hydrolysis and are therefore described as perhydrolases [30,
31]. Protein engineering has also been used to turn hydrolases into
perhydrolases [32-34].
The lipase-mediated synthesis of peroxy acids from carboxylic
acids and hydrogen peroxide can be used to perform in situ
epoxidation of alkenes [28, 35]. When unsaturated fatty acids or
their esters are treated with hydrogen peroxide in the presence of
an enzyme such as Novozyme 435, epoxidized derivatives are produced
[36]. The reaction proceeds in two steps (figure 2). Firstly,
unsaturated fatty acids are converted into unsaturated peroxy acids
by lipase-catalyzed perhydrolysis. Unsaturated peroxy or carboxylic
acids are then epoxidized via an uncatalyzed Prileshajev reaction
that is often referred to as “self-epoxidation reaction” in spite
of the fact that it proceeds predominantly via an intermolecular
process [36].
More recently, a lipase from Pseudomonas sp. was shown to
catalyze the epoxidation of cholesterol and stigmasterol in the
presence of high hydrogen peroxide concentration (50%) and ethyl
acetate, while olefins were converted into diols in the reaction
conditions tested [37].
In addition to the production of partially or completely
epoxidized free fatty acids, the chemo-enzymatic method can also be
applied to produce epoxidized plant oils [6, 38, 39]. If a
triglycerol embodying unsaturated fatty acids is treated with
H2O2 in the presence of a suitable lipase,
peroxy fatty acids are formed that epoxidize the C=C double bonds.
The resulting mixture contains epoxidized triglycerides, a small
amount of epoxidized free fatty acids, epoxidized mono- and
diglycerides. The separation of these mono- and diglycerides from
the reaction medium is difficult. Although there may be
applications where these by-products do not matter, a way to
prevent their formation has been found by adding free fatty acids
to the starting material [38]. In this way, perhydrolysis occurs
but all the hydroxyl groups of glycerol are reesterified by the
excess of free fatty acids. The resulting product only consists of
epoxidized triglycerides and epoxidized free fatty acids that can
be easily removed by alkaline washing if necessary. Using this
method, rapeseed, sunflower, soybean and linseed oil were
epoxidized with conversions and selectivities well above 90%
[6].
Chemo-enzymatic epoxidation is of considerable interest because
this method occurs in mild conditions and suppresses undesirable
ring opening of the epoxide. The epoxidation processes described by
Rüsch et al. [6] have already been carried out on the kilogram
scale. If further successful, the lipase-catalyzed perhydrolysis
could replace the problematic chemical Prileshajev epoxidation on
industrial scale.
Molecular approach of lipase-catalyzed perhydrolysis
The crystal structure of Candida antarctica lipase B shows that
this enzyme has a Ser-His-Asp catalytic triad in its active site.
The structure appears to be in an “open” conformation with a rather
restricted entrance to the active site that explains the substrate
specificity and the high degree of stereospecificity of this
lipase. In addition, the low hydrolytic activity towards
triglycerides of long chain fatty acids is also probably due to the
narrow and deep active site of the enzyme [40]. Cofactor-free
haloperoxidases, also called perhydrolases, contain a Ser-His-Asp
catalytic triad analogous to that reported for lipases [41]. Both
groups of enzymes seem to share a common origin and an analogous
catalytic mechanism, in which the involvement of the catalytic
triad would be crucial [41, 42]. Perhydrolysis presumably occurs
with an esterase-like mechanism: a carboxylic acid first reacts
with the active site serine group to form an acyl-enzyme
intermediate, which reacts with hydrogen peroxide to form a peroxy
acid. Nevertheless the Ser-His-Asp catalytic triad is not the only
determinant for perhydrolase activity because some serine
hydrolases don’t have this activity [29]. An alternate mechanism
has been proposed for perhydrolysis in which the catalytic serine
stabilizes the carboxylic acid substrate with a hydrogen bond
instead of forming an acyl-enzyme intermediate [43]. However there
is currently no experimental evidence for this proposal. An
explanation proposed to elucidate the difference in activities
between hydrolases and perhydrolases concerns the electronegative
microenvironment of the active site. The more hydrophobic
environment present in perhydrolases compared to other hydrolases
would protect the peroxy acid against hydrolysis [44]. Besides, the
presence in the structure of the enzyme of amino acids particularly
sensitive to oxidation by H2O2 and by peroxy
acids can also explain the difference in enzymatic activities [41].
Recently, Bernhardt et al. [29] performed the alignment of the
amino acid sequences of six hydrolases and six perhydrolases in
order to observe which residues appeared in perhydrolases but not
in esterases. The differing amino acids within a sphere of 12Å
around the reactive hydroxyl of the catalytic serine were mutated
by molecular biology techniques. The substitution of a single amino
acid was sufficient to shift the hydrolase activity of an aryl
esterase from Pseudomonas fluorescens to make perhydrolysis the
preferred reaction in aqueous solution. A molecular basis for the
increase in perhydrolase activity is the presence of a hydrogen
bond formed between a carbonyl oxygen atom of the enzyme and the
peroxide nucleophile. This peroxide hydroxy-carbonyl hydrogen bond
stabilizes the hydrogen peroxide attack on the putative acyl-enzyme
intermediate, hence facilitating the perhydrolysis reaction
[29].
Reaction parameters influencing the chemo-enzymatic
epoxidation
The applicability of chemo-enzymatic epoxidation using the lipase B
from Candida antarctica has been demonstrated with different
substrates such as several fatty acids, fatty acid esters including
vegetable oils, and other olefins [36, 38, 45].
Chemo-enzymatic epoxidation is influenced by various reaction
parameters. Hydrogen peroxide concentration is the most critical
parameter influencing the reaction rate and the degree of
epoxidation. Orellana-Coca et al. [46] showed that an excess of
hydrogen peroxide compared to the amount of double bonds is
necessary in order to yield a total conversion of linoleic acid
within a short time period. However, a large excess of hydrogen
peroxide results in the accumulation of peroxy acids in the final
product. These unreacted peroxy acids could be a potential problem
for reasons of safety and contamination of the final product.
Moreover, a high hydrogen peroxide concentration in the reaction
medium negatively affects enzyme activity [46]. The gradual
addition of hydrogen peroxide could be a solution to reduce the
deactivation of the biocatalyst [28, 35, 46].
Increasing reaction temperature has a positive effect on the
reaction rate of chemo-enzymatic epoxidation but it should be below
50°C to avoid hydrogen peroxide decomposition and possible enzyme
inactivation [46].
Most of the investigations have involved dilution of the
substrate in an organic solvent, particularly in toluene. Recently,
lipase-mediated epoxidation in a more environmentally-friendly,
solvent-free medium has also been reported. In these conditions,
reaction temperature has a significant impact on epoxidation. A
study concerning the chemo-enzymatic epoxidation of linoleic acid
[46] showed that the reaction in a solvent-free medium is not
complete at 30°C due to the formation of a solid or a highly
viscous oily phase, creating mass transfer limitations. Increasing
the temperature up to 60°C and using some excess of hydrogen
peroxide helped in improving the rate of epoxide formation. In
another study, the chemo-enzymatic epoxidation of oleic acid and
its methyl ester under solvent-free conditions was characterized
[47]. Epoxystearic acid and epoxystearic acid methyl ester were
synthesized with very good yields.
Stability of lipase during chemo-enzymatic epoxidation
Enzyme cost is among the important factors determining the
economics of a biocatalytic process. To make chemo-enzymatic
processes competitive with chemical processes, a high enzyme
stability and the possibility to recycle the enzyme are highly
desirable. In an investigation by Warwel et al. involving
epoxidation in a toluene containing reaction medium it was found
that Candida antarctica B lipase (Novozyme 435) was very stable,
with 75% of residual activity after 15 reaction cycles [36].
Nevertheless, when chemo-enzymatic reaction occured in a
solvent-free medium under conditions optimized for achieving high
reaction rates and epoxidized product yields, the enzyme was found
to suffer loss in activity, hence limiting its recycling [47].
Two parameters are harmful for the activity and for operational
stability of lipase B from C. antarctica (Novozyme 435) in
chemo-enzymatic epoxidation: hydrogen peroxide at high
concentrations together with elevated temperatures. Indeed, in the
presence of 6-12M hydrogen peroxide, the enzyme is rather stable at
20°C whereas at 60°C the enzyme loses activity rapidly. The rate of
deactivation increases with increasing H2O2
concentration [48]. For epoxidation processes run at elevated
temperatures, a controlled addition of H2O2
is hence important for enzyme stability. In an industrial
chemo-enzymatic process, temperature control and careful dosage of
hydrogen peroxide would be essential to optimize the enzyme
stability.
Conclusion and outlook
Despite the fact that fatty acid-epoxidizing enzymes are versatile
and useful biocatalysts, their industrial applications are still
scarce. Cytochrome P450 monooxygenases and diiron
cluster-containing monooxygenases offer an interesting possibility
for fatty acid oxidation but up to now this is an issue of academic
research only. An efficient method for the substitution of their
expensive cofactors NAD(P)H has to be found in order to use these
biocatalysts in preparative synthesis. Currently, biocatalytic
epoxidation of fatty acids using isolated enzymes is limited to the
use of lipoxygenases, peroxygenases and lipase-catalyzed
perhydrolysis followed by “self-epoxidation”. The information
provided in this review focused on the lipase-catalyzed
perhydrolysis. The recent understanding of some molecular bases of
this catalytic performance as well as the development of directed
evolution techniques, eventually in combination with rational
enzyme engineering, open up the possibility to overcome problems
yet unresolved. For example, this approach could lead to enzymes
more resistant to the drastic reaction conditions originating from
the use of hydrogen peroxide in high concentrations. Enhancing the
perhydrolysis/hydrolysis rate ratio and improving enzyme stability
to favour its recycling could reduce the cost of the process. In
this way the chemo-enzymatic epoxidation of unsaturated fatty acids
seems to be possible on industrial scale to replace the Prileshajev
chemical reaction.
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