ARTICLE
Auteur(s) : Uwe T
Bornscheuer
Department of Biotechnology and Enzyme Catalysis, Institute of
Biochemistry, Greifswald University, Felix-Hausdorff-Str. 4, 17487
Greifswald, Germany Tel.: +49 3834 86 4367, Fax: +49 3834 86
80066
Introduction
Lipases (EC 3.1.1.3, triacylglycerol hydrolases) are the most
widely used biocatalysts for the modification of fats and oils
[1-3], but they are also employed in organic synthesis [4, 5]. They
do not require cofactors, many of them are commercially available
and they exhibit high activity and stability, even in non-aqueous
systems such as organic solvents. Thus, a plethora of publications
dealing with lipases have appeared in the last decades. The
majority of examples from academic research and in industrial
applications rely on commercially available (usually immobilized)
lipases and use just a few types, which all originate from
microorganisms (i.e. lipase from Rhizomucor miehei (Lipozyme RM
IM), Thermomyces lanuginosa (Lipozyme TL IM), Candida antarctica
type B (Novozyme 435, CAL-B) or Burkholderia cepacia (Amano PS)).
In lipid modification, lipases have been often used for the
tailoring of natural lipids to meet nutritional properties,
especially for humans. The most prominent example is the synthesis
of cocoa-butter equivalent [6]. Cocoa butter is predominantly
1,3-disaturated-2-oleyl-glyceride, where palmitic, stearic and
oleic acids account for more than 95% of the total fatty acids.
Cocoa butter is crystalline and melts between 25 and 35°C providing
the desirable ’mouth feel’. Unilever [7] and Fuji Oil [8] filed the
first patents for the lipase-catalyzed synthesis of cocoa butter
equivalent from palm oil and stearic acid. Reactions are usually
performed as transesterification or acidolysis of cheap oils using
tristearin or stearic acid as acyl donors and a 1,3-specific
lipase. Structured triglycerides (sTAG) with a defined distribution
of different fatty acids along the glycerol backbone are another
area where lipases have been used, as these sTAG are important
compounds for human nutrition. sTAG containing medium chain fatty
acids at the sn1- and sn3-position and a long (preferentially
polyunsaturated) fatty acid at the sn2-position are used to treat
patient with pancreatic insufficiency and for rapid energy supply
(i.e. for sports). Another important example is Betapol™ used in
infant nutrition, which contains oleic acid at the sn1- and sn3-
and palmitic acid at the sn2-position. Currently, Betapol™ is
manufactured by interesterification of tripalmitin with oleic acid
using Lipozyme RM IM. Also, a two-step lipase-catalyzed process was
developed (figure
1), in which tripalmitin is first subjected to alcoholysis
with ethanol using a lipase from Rhizopus delemar immobilized on a
polypropylene carrier (EP-100) yielding 95% monopalmitin with a
purity >90% after crystallization. Subsequent enzymatic
esterification with oleic acid in hexane proceeded quantitatively
within a few hours and the final OPO (yield 70%) contained up to
96% palmitic acid in the sn2-position [9].
Other more recent examples for successfully industrialized
processes include lipase-catalyzed production of zero-trans
margarines (ADM/Novozymes) and diglyceride-based cooking and frying
oils (Kao Corp./ADM) [10]. The zero-trans and reduced trans oils
and fats are produced on industrial scale by transesterification
using Novozyme TL IM in combination with a cost-effective
immobilization technology.
In addition, lipases have been used on industrial case to
produce simple esters, e.g. for cosmetic applications. Prominent
examples are cetyl ricinoleate and myristyl myristate [11, 12].
Although both esters have been chemically synthesized for a long
time, enzyme technology allows higher yields and substantially
purer products. The higher costs for the biocatalyst are
compensated by energy savings (ambient temperature instead of
160-180°C) and product purification (i.e. a bleaching and
deodorization step can be omitted).
However, it often occurs that an enzyme does not meet the
requirements for a certain application and its properties have
therefore to be optimized. This usually includes the chemo-, regio-
and stereoselectivity of the biocatalyst, but also process-related
aspects such as long-term stability at high temperatures or
pH-values and activity in the presence of large substrate
concentrations need to be improved. Beside rather classical
strategies such as immobilization, additives or process
engineering, molecular biology techniques nowadays represent the
most important methodologies to tailor-design an enzyme for a given
application. Two different (but increasingly complementary)
strategies are the methods of choice: rational protein design or
directed (molecular) evolution (figure 2), which both
profited from important developments in research since the mid 90s
leading to a set of new methodologies.
This article focuses on these protein engineering methods with
special emphasis on their use to alter the properties of lipases
for lipid modification.
Background of Directed Evolution and Rational Protein
Design
Directed evolution
In principle, directed evolution is comprised of two steps: first,
the random generation of mutant libraries and second, the
identification of desired variants within these libraries using a
suitable screening or selection system. Two different strategies
for the generation of mutant libraries have been described (figure 2), an asexual
(non-recombining) evolution, in which a parent gene is subjected to
random mutagenesis to yield variants with point mutations, and the
sexual (recombining) evolution, in which several parental genes are
randomly fragmented, shuffled and reconstructed to create a pool of
recombined chimera.
In the last 15 years many methods have been developed and are
covered in a number of books [13-15] and reviews [16-18]. The most
widely used non-recombining method is error-prone polymerase chain
reaction (epPCR) [19]. Here, non-optimal reaction conditions are
used to create a mutant library [19, 20]. For example, increasing
the Mg2+ concentration, adding Mn2+ and usage
of unbalanced dNTP concentrations can substantially increase the
error rate of the commonly used polymerase from Thermus aquaticus
(Taq) from 0.001 to ~1%. The first recombining method was developed
by Stemmer (named DNA- or gene shuffling) [21] and consists of a
DNAse catalyzed degradation followed by a subsequent recombination
of the fragments without primers (self-priming PCR) and finally a
PCR with primers.
All random mutagenesis methods generate hughes mutant libraries
(usually in the range of 104-108 variants)
and rapid and highly reliable high-throughput screening or
selection systems are therefore necessary to identify desired
mutants within short experimental time. Overviews of recently
described assays can be found in a book [22] and a number of
reviews [23-27].
For libraries expressed in microorganisms, high-throughput
screening can be sometimes directly performed on colonies growing
in a solid culture like an agar-plate. Assays on agar-plated
colonies typically enable the screening of >104
variants in a matter of days, but they are often limited in
sensitivity: soluble products diffuse away from the colony and
hence only very active variants are detected or false positives
occur. Assays based on insoluble products have higher sensitivity,
but their scope is rather limited. Solid-phase screening relies on
product solubilization following an enzymatic reaction that gives
rise to a zone of clearance, a fluorescent product, a pH-shift
visualized by a pH-indicator or a strongly absorbing (chromogenic)
product like X-gal or α-naphthyl acetate and Fast Blue/Fast Red as
an example for lipase activity detection [28]. Lipolytic activity
can still be screened in a high-throughput format, on-plate, with
triolein- or tributyrin-agar through halo formation. Alternatively,
a high-throughput assay in solid phase was recently developed by
Babiak and Reymond using esters of coumarin [29].
However, many assays cannot be applied in a solid-phase format.
Thus, individual clones must be grown and assayed in microtiter
plates (MTP). These assays are significantly more time-consuming
than solid-phase assays. However, by using robot automation and
colony picking technology, throughput can be substantially
increased. In addition, MTP-based assays have the major advantage
that screening provides significantly more information compared to
a selection approach as the activity can be directly and
quantitatively measured and even allows to determine the kinetics.
For lipases, these assays are usually based on commerically
available p-nitrophenyl esters of varying fatty acid chain length,
but also the use of resorufin or umbelliferyl esters has been
described. Hydrolysis of these substrates releases a
chromophore/fluorophore that can be quantitatively measured
time-resolved at high sensitivity. However, these artificial
substrates differ from the true lipase substrate and hence can lead
to false positive hits. Furthermore, the substrates are often
unstable at extreme pH or temperature and this autohydrolysis can
limit their use in certain screening efforts. Alternative compounds
were suggested by Reymond et al. coupled with a periodate treatment
and β-elimination to release the chromophore/fluorophore [30-32].
However, these substrates have to be chemically synthesized and
only end-point measurements are possible. For the screening of
lipases active in the synthesis (i.e. esterification or
transesterification) reaction, a fluorometric method was described
as well [33]. This method is based on the transesterification
between an alcohol and a vinyl ester of a carboxylic acid.
Acetaldehyde generated from the vinyl alcohol by keto-enol
tautomerization is reacted with a (non-fluorescent) hydrazine
(NBD-H) to produce the corresponding highly fluorescent hydrazone,
which is then quantified by fluorimetric measurement (figure 3).
Alternatively, lipase synthesis activity can be indirectly
measured in an organic solvent using a modified p-nitrophenyl ester
assay [34]. Transesterification activity of immobilized esterases
was determined by sampling p-nitrophenol released and
subsequent spectrophotometric quantification in an aqueous system.
A similar method was reported very recently for the determination
of the synthesis activity of a lipase [35].
A totally different strategy to find the best variant in a
mutant library is bacterial surface display. Kolmar et al. showed
that E. coli bacteria that display esterases or lipases on
their cell surface together with horseradish peroxidase (HRP) are
capable of hydrolyzing carboxylic acid esters of biotin tyramide.
The tyramide radicals generated by the coupled lipase-peroxidase
reaction were short-lived and therefore became covalently attached
to reactive tyrosine residues that are located in close vicinity on
the surface of a bacterial cell that displayed hydrolase activity.
Differences in cellular esterase activity were found to correlate
well with the amount of biotin tyramide deposited on the cell
surface. This selective biotin tyramide labeling of cells that had
lipase activity allowed their isolation by magnetic cell sorting
[36].
Rational design
Engineering a protein by rational design requires the availability
of the enzymes tertiary structure or at least a homology model of
sufficient quality. Furthermore, detailed information about the
structure-function relationship and usually the reaction mechanism
is required to allow for the prediction of amino acid residues to
be mutated. In the past decades, the number of protein structures
deposited in the Brookhaven protein database (pdb) and protein
sequence information in various databases substantially facilitated
the rational design of proteins. Furthermore, a plethora of
modeling software has been developed, which makes this methodology
easier to use and also enhances the success rate of modeling
predictions. Usually, the information derived from computer
modeling identifies certain amino acids (hot spots) which should be
altered to lead to a change in the enzymes properties such as
broadened or restricted substrate range or altered selectivity.
Site-directed mutagenesis (SDM) is then performed at these
positions using for example, the QuikChangeTM
Site-Directed Mutagenesis method from Stratagene. In many cases, it
might be more advantageous to directly perform a saturation
mutagenesis at the selected position(s), which will introduce all
19 proteinogenic amino acids and hence increase the chance to find
desired variants. A combination of rational protein design with
directed evolution (CASTing) has been described by the Reetz group
[37].
Examples
The chain-length selectivity of lipases was altered by rational
protein design and SDM, as shown by Joerger and Haas for the
Rhizopus oryzae (formerly Rhizopus delemar) lipase (RDL) [38, 39].
Based on the crystal structure, they applied molecular modeling to
identify the molecular determinants of acyl chain length
specificity of this enzyme and were able to change this property
significantly. In another example – while trying to isolate new
enzyme variants of the extracellular lipase from Thermomyces
lanuginosa with enhanced activity in the presence of detergent –
Danielsen et al. randomized nine amino acids in two regions
flanking the flexible α-helical lid. A S83T mutation was found in
six of the seven most active variants, which in the homologous RDL
had been proven to determine the chain-length preference [40]. More
recently, an esterase was subjected to directed evolution and a
mutant was identified having a lipase-like chain-length specificity
[41]. Moreover, the variant also was shown to have distinct
sn2-specificity, a unique feature, which makes this enzyme very
attractive for the synthesis of structured triglycerides as
outlined in the introduction. A similar change in chain-length
specificity was recently achieved by random mutagenesis of a
distinct region of an esterase from Pseudomonas fluorescens. The
best variants had a 10-fold higher catalytic activity towards
p-nitrophenyl dodecanoate than the wild-type enzyme [42].
Fujii et al. reported the enhancement of amidase activity of a
Pseudomonas aeruginosa lipase after one single round of random
mutagenesis. Mutant libraries were screened for hydrolytic activity
against oleyl-naphthylamide vs. the hydrolysis of the corresponding
carboxylic acid ester. Three mutational sites were identified to
enhance amidase activity, and the double mutant F207S/A213D was
found to have the highest amidase activity, 2-fold that of the
wild-type. These mutations were located near the calcium binding
site, far from the active site [43]. For organic synthesis, the
stereoselectivity of lipases is one of the most important features
and directed evolution was also applied to alter this
characteristics. Thus, Reetz et al. enhanced the selectivity of a
lipase from Pseudomonas aeruginosa for the kinetic resolution of
2-methyldecanoic acid p-nitrophenyl ester from E=1.1 to the
practically useful value of E=51 [44, 45].
CAL-B is probably the most useful lipase and it could be show
that its thermostability could be improved by directed evolution as
variants were found after two rounds of epPCR, that are 20-fold
more stable at 70°C than the wild type. Positions 221 and 281 were
found to be critical to prevent irreversible inactivation and
protein aggregation and the variants were also found to be more
active against p-nitrophenyl butyrate and
6,8-difluoro-4-methylumbelliferyl octanoate [46].
Stability of lipases is very important in large scale
applications to make processes more cost-efficient. In order to
enhance the stability of a lipase from Rhizopus oryzae (ROL)
towards lipid oxidation, products such as aldehydes, six lysine and
all histidine residues (except for the catalytic His) out of 22
amino acid residues (15 Lys, 7 His, figure 4) prone to react
with aldehydes were chosen. These selected positions were then
subjected to saturation mutagenesis using the gene encoding the
prolipase. In order to quickly and reliably identify stability
mutants within the resulting libraries, active variants were
pre-screened by an activity staining method on agar plates. Active
mutants were expressed in E. coli in a 96-well MTP format and
a stability test using octanal as model deactivating agent was
performed. The most stable histidine mutant (H201S) conferred a
stability increase of 60%, which was further enhanced to 100% by
combination with a lysine mutant (H201S/K168I). This increase in
stability was also confirmed for other aldehydes. Interestingly,
the mutations did not affect specific activity, as this was still
similar to the wild type enzyme [47].
Further examples for the successful application of protein
engineering of lipases and esterases are summarized in table 1.
Table 1 Selected examples of lipases (and esterases)
improved by protein engineering methods.
|
Enzyme (origin)
|
Target
|
Mutagenesis method
|
Assay
|
Improved property
|
References
|
|
P. aeruginosa lipase
|
Enantioselectivity
|
epPCR and others
|
- MTP assay with chiral
- p-nitrophenyl esters
|
Increased enantioselectivity
|
[44, 45]
|
|
P. aeruginosa lipase
|
Improve amidase activity
|
- epPCR
- Saturation mutagenesis
|
Activity staining on agar plate
|
2-fold higher amidase activity
|
[43]
|
|
P. aeruginosa lipase
|
Substrate specificity
|
CASTing
|
MTP assay with p-nitrophenyl esters
|
Expanded substrate acceptance for different carboxylic acid
esters
|
[37]
|
|
R. oryzae lipase
|
Substrate specificity
|
Saturation mutagenesis
|
Agar-plates containing rhodamine B
|
Altered chain-length preference
|
[38, 39]
|
|
R. oryzae lipase
|
Enhanced stability
|
Saturation mutagenesis
|
MTP assay
|
Increased stability towards aldehydes
|
[47, 48]
|
|
R. arrhizus lipase
|
Thermostability
|
|
Agar-plates containing rhodamine B
|
Improved thermostability and higher temperature optimum
|
[49]
|
|
C. antartica lipase B
|
Thermostability
|
- epPCR
- Saturation mutagenesis
|
MTP with p-nitrophenyl esters and heating
|
>20-fold improvement in half-life at 70°C
|
[46]
|
|
B. gladioli esterase
|
Stability in organic solvents
|
epPCR
|
pH indicator
|
100-fold improvement of activity in 35% DMF
|
[50]
|
|
B. subtilis lipase A
|
Enantioselectivity towards 1,2-O-isopropylidene-glycerol
|
Saturation mutagenesis near the active site
|
Phage display
|
Inverted enantioselectivity
|
[51]
|
|
Esterase from deep sea
|
Broadened substrate range
|
epPCR
|
MTP with p-nitrophenyl esters
|
Lipase-like chain-length selectivity, also sn2-specific
|
[41]
|
|
P. fluorescens esterase
|
Broadened substrate range
|
Randomized saturation mutagenesis
|
MTP with p-nitrophenyl esters
|
Lipase-like chain-length selectivity
|
[42]
|
Conclusion
Protein engineering methods have emerged as very powerful tools to
alter the properties of enzymes for biocatalysis. Especially
directed evolution became a mature technology within just a few
years and the diverse set of molecular biology tools to create
well-balanced mutant libraries as well as suitable high-througput
screening methods allowed to create biocatalysts including lipases
with substantially altered properties. Furthermore, rational
protein design was shown to be a useful method to change lipase
specificities and with an increasing number of protein structures
available, this approach will certainly become even more important
in the future.
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