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3
Directed Enzyme Evolution
and High-Throughput Screening
Michael J. McLachlan,1 Ryan P. Sullivan2 and Huimin Zhao3
1
Center for Biophysics and Computational Biology, University of Illinois
at Urbana-Champaign, Urbana, IL 61801, USA
2
Department of Chemical and Biomolecular Engineering, University of Illinois
at Urbana-Champaign, Urbana, IL 61801, USA
3
Departments of Chemical and Biomolecular Engineering, and Chemistry, University
of Illinois at Urbana-Champaign, Urbana, IL 61801, USA
3.1 Introduction
Early enzyme products developed for industrial applications were produced by native hosts
via fermentation and consisted of a complex mixture of secreted enzymes produced at low
yields. Now, over 90% of industrial enzymes are produced recombinantly for maximal purity
and productivity [1]. The integration of enzymes into a commercial process relies on the
availability of those with high activity and stability under process conditions, desired
substrate specificity, and high selectivity. More often than not, naturally occurring enzymes
do not fulfill the requirements of these harsh industrial conditions, and optimization is
necessary to obtain a suitable enzyme catalyst for production needs. This tailoring of
enzymes can be accomplished through two experimental routes. The first is rational design,
which targets specific residues of a protein for mutagenesis to predetermined amino acid
mutations, and is only applicable when there is detailed knowledge of the relationships
between the enzyme’s structure and mechanism/function. And while an increasing number of
enzymes are being characterized, the majority do not have this depth of information readily
available, as it requires considerable effort to obtain. In the absence of this information, the
tailoring of an enzyme can still be accomplished through the second route: directed
evolution.
Biocatalysis for the Pharmaceutical Industry : Discovery, Development, and Manufacturing edited by J. Tao, G.-Q. Lin, and A. L.
© 2009 John Wiley & Sons Asia (Pte) Ltd. ISBN: 978-0-470-82314-9
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Directed evolution is the general term applied to the combined techniques of generation
of a library of protein mutants (or variants) and selection of a protein with desirable
function from within that library [2]. It is an iterative Darwinian optimization process,
whereby the fittest variants are selected from an ensemble of mutants [3]. Directed
evolution can be used to target a number of enzymatic characteristics, including activity,
substrate specificity, thermal and oxidative stability, enantioselectivity or enantiospecificity, pH optima or range, and tolerance to solvent [4]. While a typical directed evolution
experiment focuses on a single enzymatic trait, there are some examples of improving
several traits simultaneously.
Developed primarily in academic laboratories, directed evolution practices in industry
are still in their infancy and present an imposing challenge for process scientists today.
Choosing the appropriate methods of library generation and screening or selection is
paramount to the success of any directed evolution experiment. Library diversity can be
created through either mutagenesis (random or semi-rational) or gene recombination, and
which of these methods is chosen depends on many factors, such as the availabilities of
homologous genes, structural knowledge, and characteristic data of the enzyme of interest.
The library size created is typically very large ( > 104–6), and close evaluation of each
variant is not feasible. The need for a method to find the improved ‘needle in a haystack’
enzyme becomes evident, with several strategies, including selection, enrichment, and
high-throughput screening, offering ways to sift through the library clutter and find a
variant with the desired enzymatic trait. However, once an evolved enzyme has been found
that exhibits improved characteristics, the artificial conditions in which the selection
method was carried out may result in an enzyme whose properties may not carry over to the
real biocatalytic process. Therefore, the more similar a screening system is to the actual
application process, the more likely it is to find an improved enzyme that will be
complementary to the application.
The aims of this chapter are to familiarize the reader with the various techniques used in
library creation and high-throughput screening, as well as representative examples in the
pharmaceutical industry and the technology used to adapt the laboratory-based techniques to
industrial settings. The topics covered do not represent a complete listing of methods utilized,
as more strategies for library creation and high-throughput screening are constantly
introduced.
3.2 Directed Evolution Library Creation Strategies
Library creation strategies generally fall into three main categories: random mutagenesis,
semi-rational design, and gene shuffling (Figure 3.1). Random mutagenesis introduces
mutations throughout a target gene encoding for the industrial enzyme of interest. These
mutations may be in the form of point mutations (either transitions or transversions), insertions,
deletions, inversions, or frame-shift mutations. Semi-rational design is a combination of
random mutagenesis and site-directed mutagenesis, in which specific residue positions are
rationally determined to play important roles in the enzyme’s function, and are subsequently
randomized to all 20 amino acids. Gene shuffling involves exchanging fragments of genes with
one another to create a library of chimeric progeny. In directed evolution studies, this is
typically accomplished by homologous recombination for sequences with high similarity or by
nonhomologous recombination for those with low similarity.
Directed Enzyme Evolution and High-Throughput Screening
47
Figure 3.1 Overview of DNA library creation strategies. Random mutagenesis introduces mutations at
positions throughout the gene sequence. Semi-rational design randomizes only the specific position(s) of
interest. Gene shuffling brings existing sequence diversity from different parental DNA sequences
together to form a chimeric library
3.2.1 Random and Semi-Rational Mutagenesis
The most commonly used method for random mutagenesis library creation is error-prone
polymerase chain reaction (epPCR), which introduces mutations in a gene product by lowering
the fidelity of DNA polymerase. This can be achieved through many different reaction
conditions, including use of nucleotide base analogs and alkylating agents into the polymerase
chain reaction (PCR) reaction mixture, but severe biases typically result from such modification. The more favored protocols utilize an Mg2þ -dependent polymerase that lacks exonuclease activity, substitute Mn2þ for Mg2þ , and extend DNA products with an uneven mixture of
the four precursor deoxynucleotidetriphosphates [5]. The mutation rate can be easily adjusted
by varying the concentration of Mn2þ in the reaction mixture or by changing the number of
cycles of amplification. Typically, a library is created with a low mutation rate (one to five base
pairs, or one to three amino acids per gene) to prevent disruption of enzymatic activity and
generation of a library too large to screen comprehensively. However, where there is an efficient
selection scheme or robust high-throughput screen available, higher mutation rates have been
successful in gaining improved function [6,7].
epPCR is considered as a random mutagenesis technique, but the term random should be
interpreted loosely. Ideally, a protein library should satisfy the requirement of having an equal
probability of substitution of any of the 20 amino acids randomly into any residue position of
the protein. However, the library creation is performed at the level of the gene encoding the
protein; thus, an ideal distribution is improbable due to the redundancy of the genetic code.
Mutations at the wobble position (third base of the codon) typically result in a neutral mutation,
and the frequency of two adjacent bases being mutated is considerably low, which results in
only six possible amino acids on average at each position. As a result, the DNA sequences of
selected progeny with improved function are determined, and positions at which mutations
have been incorporated are sometimes subjected to an additional round of screening using
saturation mutagenesis. Saturation mutagenesis involves creating a library by designing
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degenerate primers for the residue position in question and screening the resultant library to
determine which of the 20 amino acids results in the most dramatic improved effect at that
position. Gene site saturation mutagenesis (GSSM) takes this concept to its fullest extent by
creating a library consisting of all 20 possible amino acid substitutions for each position in a
protein [8,9].
Sequence saturation mutagenesis (SeSaM) [10] is an alternative four-step approach to
epPCR that overcomes several barriers faced by its predecessor. First, a library is created
with standard nucleotides with the presence of a-phosphothionate nucleotide, an alkaline
labile analog. The incorporated analog is hydrolyzed, thus creating a library of fragments of
random length. The fragments are then treated with a terminal deoxynucleotidyl tranferase
(TdT) to incorporate a random number of universal bases at the 30 -termini, followed by
elongation of labeled gene product to full length. A concluding PCR then replaces the
universal base substitutions with standard nucleotides. The mutations incorporated into the
SeSaM library are randomly distributed throughout the parent gene template and independent of the DNA polymerase, thus avoiding any potential mutational bias and resulting in
the exchange of an amino acid at any position to all other 19 possible amino acids. The
SeSaM protocol has also been modified to enrich the transversion rate and allow for
adjustable mutational biases [11]. However, the use of different base analogs, biotinylated
primers, and the additional time-consuming steps for library creation are drawbacks to this
method.
Insertion and deletion mutagenesis expands the reach of random mutagenesis to include
alteration of the size of the gene of interest when generating a library. This process requires the
use of transposable elements that insert randomly into the target and are subsequently excised,
leaving behind an in-frame fragment. Pentapeptide scanning mutagenesis results in the
insertion of a 15-bp fragment, but the amino acids incorporated are predetermined due to
the cognate target site of the transposon and preset restriction site sequence [12]. This results in
a less than optimal library with less diversity and more bias than other methods. It is particularly
suited towards obtaining mutants whose loop structures dictate activity enhancements. An
improved technique of the same style is random insertion/deletion, which makes it possible to
delete a multitude of consecutive bases from random locations in a gene and insert either a
specific or randomized sequence of arbitrary length in its place [13]. The diversity obtained
from this method far outstretches that of conventional epPCR, but it comes at the cost of several
additional processing steps.
3.2.2 Gene Shuffling
Although random mutagenesis techniques offer a relatively simple scheme to approach a
desired enzymatic trait, the frequency of gaining beneficial mutations is generally low.
Typically, only one or two amino acid changes are generated in each round, with higher
random mutation rates leading to either a library too large to screen successfully or to
incorporation of mutations that lead to loss of function. Gene shuffling can overcome this
limitation by allowing a large number of beneficial mutations from multiple genes to
incorporate at a single step of the library creation process. There are a variety of shuffling
methods available, which can be divided into categories of homology dependent and homology
independent.
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3.2.2.1 Homology-Dependent, Primer-Independent Assembly
The pioneering technique of DNA shuffling [14] was a breakthrough for in vitro recombinant
mutagenesis and diversity generation. The method is centered on taking a number of
homologous parental template genes, randomly digesting them with DNase I and reassembling
the pieces back together into full-length genes through a primerless PCR reaction (Figure 3.2a).
In this extension process, pieces of different genes can anneal to each other at homologous
regions and extend to full-length fragments through the aid of DNA polymerase. DNA shuffling
can be applied to combining beneficial mutations found from random mutagenesis screening,
which can reduce the processing steps required to determine which mutations screened for are
actually necessary for improved function. A single gene variation of DNA shuffling involves
incorporation of mutations at the extension level, screening for improved progeny, and
subsequent shuffling of these mutants for further screening. In contrast, family shuffling [15]
begins the process with naturally occurring homologous genes (typically > 60% identical).
Because the parental templates in this process have already been subjected to natural selection,
much larger numbers of mutations are tolerated, leading to a library with a broader sequence
space that maintains a higher percentage of functional enzymes.
These methods all have similar limitations, however, as they rely on sequence homology for
recombination events to occur. Gene fragments will tend to anneal to each other only at areas of
Figure 3.2 Examples of gene shuffling methods used for DNA library creation in directed evolution.
(a) Homology-dependent primer-independent DNA shuffling; (b) homology-dependent primerdependent StEP; (c) homology-independent SHIPREC
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high sequence similarity, leaving out the possibility of crossovers in other regions with
low similarity. Also, the chimeric genes typically only contain one to four crossover
events, which strongly limits the accessible sequence space. This low crossover rate of DNA
shuffling was addressed with a method called random chimeragenesis on transient templates
(RACHITT) [16]. The parental template in this case contained uracil and was single stranded
to serve as a scaffold for hybridization of second-strand fragments from homologous genes.
The crossover rate was dramatically increased (around 14 per chimera product), leading to
much greater diversity in the library. However, this method is more labor intensive and is
subject to optimization of reaction times and temperatures to obtain full-length genes for
screening.
3.2.2.2 Homology-Dependent, Primer-Dependent Assembly
In contrast to the methods described above, several techniques have been developed that
require end-primers or addition of oligonucleotides to encourage frequent crossover between
templates. Random priming recombination [17] involves a parental template gene as a scaffold,
with random primers used in place of DNase I fragments to anneal randomly on the template
and extend to full-length genes. A staggered extension process (StEP, [18]) starts the extension
step of shuffling only with end-primers. Instead of extending the gene to full length in one cycle,
the process is carried out through very short extension and annealing steps, resulting in
template switching throughout the course of full-length gene assembly, thereby producing
multiple crossovers (Figure 3.2b). However, the limitation in combining close-proximity
mutations is still present. Synthetic shuffling is accomplished without any full-length gene
templates, but rather with synthetic degenerate oligonucleotides designed with bioinformatic
information on the gene of interest [19]. The oligonucleotides contain overlapping ends that
anneal and extend into full-length composite genes. This method opens up the capabilities of
introducing site-directed mutations (via semi-synthetic shuffling) or codon optimization into
the gene library. Biased mutation-assembling [20] takes wild-type and mutant genes with
distinct beneficial mutations, defines blocks within them containing only single mutations, and
amplifies the regions with small overlaps to adjacent blocks. The blocks are then reassembled
by overlap extension PCR, but are combined in ratios which favor incorporation of mutations
which displayed the most improvement.
3.2.2.3 In Vivo Assembly
All of the gene shuffling methods discussed so far involve the application of in vitro techniques
for generation of the gene library. Another possibility to create low-complexity chimeric
libraries is through in vivo recombination methods. In combinatorial libraries enhanced by
recombination in yeast [21], parental genes are again randomly fragmented with DNase I and
reassembled in a primerless reaction. However, the annealing step is repeated at lower and
lower temperatures as the reaction progresses (‘progressive hybridization’), facilitating the
annealing of low-homology genes. Amplification of full-length genes is then carried out with
primers designed with overlapping regions on the expression vector. The linearized vector and
library of genes are then transformed into yeast to promote in vivo homologous recombination.
Because there is typically more than one gene hybrid transformed into a yeast cell, recombination results in an increased diversity.
Directed Enzyme Evolution and High-Throughput Screening
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Heteroduplex recombination involves preparing single-stranded DNA from two different
homologous genes and mixing them to form heteroduplexes. These heteroduplexes are then
transformed into a host that then creates hybrid homoduplexes through in vivo mismatch repair
mechanisms [22].
Another technique takes advantage of recA-mediated homologous recombination in a recBC
sbcA Escherichia coli mutant [23]. In this method, the parental genes flank a linearized plasmid,
and transformation along with in vivo homologous recombination results in single crossover
progeny on circularized plasmids. Repeating this process produces one crossover per iteration,
and can be modified to include a different parental gene per round.
3.2.2.4 Homology-Independent Assembly
As stated earlier, one of the major drawbacks to DNA shuffling and techniques derived from
this method is that crossovers occur only at homologous regions. If low-homology parent genes
are used in these methods, then the majority of product genes tend to be parental themselves
instead of hybrids [24]. In an attempt to dissociate the creation of hybrid enzymes from DNA
sequence homology, a technique called incremental truncation for the creation of hybrid
enzymes (ITCHY) was created [25]. In this procedure, two parent enzymes are incrementally
truncated with exonuclease III under controlled conditions, and the various generated
50 -fragments and 30 -fragments are ligated back together to form a library of chimeric
sequences. The resulting enzymes screened are fusions of the amino-terminal portion of one
parent and the carboxy-terminal end of the other parent. However, careful optimization and
control of the fragment generation is required for a successful library, and makes the method
difficult and time consuming. In addition, the fusion enzymes created contain only one
crossover region, are not necessarily full-length genes, and can be conjoined at places that are
not at structurally related sites [26].
A modified version of this protocol, called THIO-ITCHY, includes the random incorporation
of a-phosphothioate nucleotide analogs into the parent genes [27,28]. Exonuclease III activity
is inhibited at sites of analog incorporation, which relieves the efforts of producing incremental
truncation aliquots. In combination with epPCR, the diversity of the fusion libraries created can
be further expanded.
Yet another variant of the ITCHY method, fittingly called SCRATCHY, was developed
through the combination of ITCHY with DNA shuffling to create multiple crossover libraries
independent of homology [27,28]. Further development of this technique produced enhancedcrossover SCRATCHY, in which amplification of ITCHY hybrids is carried out in defined
blocks with skewed primers (a forward primer from one parent and a reverse primer from the
other). These amplified fragments are then pooled and subjected to DNA shuffling, resulting in
a process that selectively enriches hybrids that contain multiple crossovers [29].
Another method that produces hybrid genes independent of homology is sequence homology independent protein recombination (SHIPREC) [30]. Parental genes are first fused together
via a linker containing a unique restriction site to form heterodimers. These heterodimers are
then randomly digested with DNase I in a controlled reaction. The fragments corresponding to
the length of either parental gene are then isolated, blunt-end digested, and ligated to create
circularized gene hybrids. These hybrids are then linearized with the unique restriction site
introduced originally into the linker (Figure 3.2c). The library is still limited to one crossover
event, but is also more likely to contain structurally conserved crossover events.
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3.2.2.5 Multiple-Parent, Nonhomologous Assembly
All of the previous homology-independent shuffling methods are limited by the use of only two
parental genes. However, methods to recombine DNA from multiple parents do exist. In cases
where gene templates are from eukaryotic sources, exon shuffling can be implemented to
interchange domain-encoding exons [31]. In this way, functional domains are connected
through design of oligonucleotides used to amplify permitted crossovers, and the random
aspect of assembly is avoided. Random multi-recombinant PCR (RM-PCR) is a more
generalized scheme of exon shuffling in which reassembly of blocks into full-length genes
is possible through overlap-extension PCR, with oligonucleotides containing the crossovers,
and can be used for prokaryotic and eukaryotic genes alike [32].
Degenerate homoduplex recombination is similar to synthetic shuffling, and utilizes a set of
degenerate top-strand primers that are designed to encompass the diversity information of the
genes to be shuffled. Gaps in between these top-strand primers are filled in using dephosphorylated bottom-strand oligonucleotides as templates [33].
Nonhomologous random recombination assembles DNase I-digested fragments that have
been blunt-end polished prior to ligation. The addition of hairpin oligonucleotides that cap only
the ends of the full-length genes results in preferential ligation of intermolecular pieces [34]. To
address limitations with creating frame-shift mutants with premature stop codons, the addition
of a chloroamphenicol acetyltransferase (CAT) fusion allowed for preferential selection
against truncated or insoluble proteins [35].
There is no single library creation method that will satisfy the ideal requirements of highdiversity quality, large library size, and ultimately the feasibility of comprehensive screening of
the generated library. Evaluation of each case is necessary to determine which library creation
method is suitable for the project’s goals.
3.3 Directed Evolution Library Screening/Selection Methods
The options for analyzing a library will be guided in part by the properties of the protein being
studied. Whether the target is a binding protein or an enzyme, what substrates or derivatives are
available and how these are linked to metabolism will help determine the appropriate screening
technique. The analysis involves finding a method that will link the desired activity with the
gene variant that encodes it, and can be split into two approaches: screening and selection. In a
screening method, variants in the library are assayed individually. For example, by measuring
the enzymatic activity in a cell lysate, both active and inactive clones will be discovered.
Alternatively, selection methods deal with the entire library at once. For example, by applying
antibiotics to bacteria on an agar plate, only resistant clones will grow. A selection method is
desirable, since it increases the size of the library that can be practically assayed. The methods
used can also be classified as either in vivo (where intact cells are used) or in vitro (where
isolated cellular components are utilized). The following will give an overview of these
methods, with more details available in published reviews [36,37].
3.3.1 In Vivo Methods: Genetic Complementation
The classic microbiological approach of genetic complementation can be very useful as a
directed evolution selection [38,39]. The activity being investigated is intrinsically linked to
Directed Enzyme Evolution and High-Throughput Screening
53
the growth of a microbe such as E. coli or Saccharomyces cerevisiae, as it either provides an
advantage to the wild-type strain or the library can complement a particular mutant strain. By
creating an E. coli strain that was unable to utilize glucose as a carbon source, an overexpression
library was screened to discover genes with latent glucokinase activity [40].
When the desired reaction is not essential, creative thinking must be employed. Adding a
selectable moiety may create derivatives of a natural substrate that allow for selection.
Hwang et al. [41] used this approach to derive a screen for enantioselective hydrolases. By
linking the antibiotic chloramphenicol to either the R- or S-enantiomer of 2-phenylbutyric
acid, they showed that Exiguobacterium acetylicum could hydrolyse the R-form (as the
released chloramphenicol inhibited growth) but not the S-form. However, one must be aware
that activity on a derivative may not always correspond to the activity on the actual desired
substrate.
The degree of selection can be adjusted as needed by the use of different vectors or promoters
to alter the expression level of the enzyme within the cell. Alternatively, the stringency of
selection can be increased by channeling the substrate away by adding a competing
pathway [42].
3.3.2 In Vivo Methods: Chemical Complementation
The drawback of traditional genetic complementation approaches is that they are specific for
the gene being studied, and new screens must be devised for each enzyme being investigated.
Chemical complementation methods aim to be more general in their approach, by using the
substrate of the targeted reaction to influence an unrelated reporter system. This allows one to
use well-studied reporters, such as b-galactosidase, or the amino-acid-selectable markers of
yeast if they can be linked in some manner to the substrate.
This has been accomplished through the use of the yeast three-hybrid assay [43]. Here, the
phenotypic readout is the blue color produced by the action of b-galactosidase on X-gal or onitrophenol-b-D-galactopyranoside. The transcription of this reporter gene is determined by
the dimerization of a methotrexate-binding DNA binding domain to a dexamethasone-binding
activation domain. The two components are linked by a composite molecule containing the
substrate of interest, methotrexate, and dexamethasone. If the library variant cleaves the
substrate, then there is no dimerization of the transcriptional components, b-galactosidase will
not be produced, and there will be no blue color. By changing the reporter gene, this type of
screen could be converted to a selection, as was demonstrated with a glycosynthase, where
bond formation controlled the transcription of a leucine biosynthetic reporter [44].
Other approaches could use transcriptional regulators that directly bind the substrate or
product of the reaction and activate the reporter gene. For instance, a mutant transcriptional
activator from Pseudomonas putida, NahH, was used that can bind various benzoic acids to
develop a screening/selection method to detect the action of benzaldehyde dehydrogenase [45].
A transcriptional regulator may need to be engineered to bind the desired compound before it
can be used in such a manner [46].
3.3.3 In Vivo Methods: Surface Display
The ability to display proteins on the surface of an organism has been exploited as a screening
method, typically in conjunction with fluorescence-activated cell sorting (FACS) [37,47,48]
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Figure 3.3 Cell surface display. Proteins displayed on the cell surface of organisms bind a fluorescent
molecule. Cells are passed through a FACS machine, allowing separation into populations that do or do
not bind the target
(Figure 3.3). The approach is particularly well suited for engineering binding proteins such as
antibodies, although it has also been applied to enzymes. One of the first widely used surface
display methods was through the use of phage [49]. By inserting a target protein into one of the
coat proteins of filamentous phage (initially protein pIII, but others such as pVIII have also been
used), very large libraries (1010) of variants can be expressed in E. coli and subsequently
packaged into active phage. Antibodies are used to enrich the desired variants in vitro, and the
phage can then reinfect bacteria in multiple rounds of selection. The selection strategy is useful
for increasing the binding affinity of a protein. Stability can also be engineered through the use
of proteases, which will cleave a poorly folded protein more rapidly and reduce the infectivity
of the phage [50].
Another method is yeast display, which consists of fusing the engineering target to the
C-terminus of the Aga2 cell surface agglutinin protein, allowing exposure to fluorescently
labeled substrates in the media [51]. Bacterial display systems have also been produced using a
number of different scaffold proteins, such as the outer membrane proteins OmpA or OmpX,
the thioredoxin protein within the bacterial flagella, or autotransporters such as AIDA-I [37].
Proteins may express differently depending on which scaffold they are fused with, or whether
bacteria or yeast are used. The use of yeast as an expression host may be beneficial if the protein
requires post-translational modifications.
Directed Enzyme Evolution and High-Throughput Screening
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3.3.4 In Vitro Methods: Lysate Assay
Perhaps the most direct way of analyzing a library is to perform the functional assay on the
protein itself. Individual clones from cells transformed with the library can be grown and
made to express the protein of interest. This can then be isolated and the relevant assay
performed. A cell lysate will often contain enough of the protein such that further purification
is not necessary for screening purposes. Using microplates increases the throughput of the
screening and is useful where the progression of a reaction can be monitored visually, either
by the distinct absorbance of the substrate or product or of a coupled reaction. The throughput
of this type of screen is comparatively low, on the order of 104, but the approach is flexible and
easily implemented. Reaction products from such assays could also be detected directly
through the use of gas chromatography, high-performance liquid chromatography, or mass
spectrometry.
3.3.5 In Vitro Methods: Ribosome Display
Ribosome display is an in vitro display method by which one can link the protein of interest to
the gene coding for it. As demonstrated with a single-chain scFv antibody, by using in vitro
transcription and translation on DNA lacking a stop codon, a complex consisting of the
mRNA, ribosome, and translated protein is generated [52]. The variants of interest are then
selected by exposing the complex to the protein’s binding partner and washing away
nonbinding complexes. The bound complexes are dissociated and the mRNA molecules
can be collected and converted back into DNA via reverse transcription, resulting in a pool of
sequences enriched for the desired binding activity. A similar approach is used in the
technique of mRNA display, but the protein of interest is linked to its mRNA by the use of a
puromycin molecule [53,54]. Both techniques have the advantage of being completely
in vitro, increasing the potential size of the library (to 1013) and the speed of screening by
avoiding transformation steps into cells.
3.3.6 In Vitro Methods: In Vitro Compartmentalization
Inspired by the natural linkage of genotype and phenotype observed with cells, Tawfik and
Griffiths [55] introduced the idea of in vitro compartmentalization (IVC) by coupling
transcription and translation within a water-in-oil emulsion. By forming aqueous droplets of
around 2 mm in diameter, genes, their proteins, and more importently substrates and products of
the protein are all contained, which allows for the selection of catalytic activities. The concept
was initially demonstrated on enzymes that act on DNA, such as HaeIII methyltransferase,
where the catalytic activity acted directly on the genotype. The use of antibodies and
streptavidin-coated microbeads allowed attachment of a gene, its encoded protein, and biotintagged substrate which was converted to product. An antibody to the product allowed the
isolation of improved variants of the enzyme phosphotriesterase through flow cytometry [56].
Extending the emulsion to a water-in-oil-in-water mixture allowed further refinement of the
IVC concept. Compartmentalization of E. coli containing serum paraoxonase variants allowed
the accumulation of fluorescent product to a point where it could be detected by FACS [57]. This
approach was also used with in vitro transcription and translation to evolve b-galactosidase
activity from the Ebg gene [58].
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3.3.7 Equipment/Automation
Directed evolution relies on the analysis of large numbers of clones to enable the discovery of
rare variants with improved function. In order to analyze these large libraries, methods of
screening or selection have been developed, many of which use specialized equipment or
automation. These range from the use of multichannel pipettes, all the way up to robotics,
depending on the level of investment [59]. Specialized robotic systems are available to perform
tasks such as colony picking, cell culture, protein purification, and cell-based assays.
A very versatile piece of equipment that is affordable for individual laboratories is the
microplate reader. This allows multiple samples to be analyzed at once, commonly in a 96-well
format, although 384- and 1536-well formats are available. Typical measurements that can be
performed include UV–Vis absorbance, fluorescence, or luminescence, allowing a range of
assays to be performed, such as cell growth, enzyme kinetics, enzyme stability, or enzymelinked immunosorbent assay [60–62]. Functionality can be increased by the use of liquid
dispensing systems or automatic plate handling.
Flow cytometry is increasingly used in high-throughput screening [63]. Here, a cell solution
is pressurized into a stream and directed through various lasers, with information on forward
and side scatter and on fluorescence being collected. Based on this data, the cells can be
directed into separate pools. The throughput of these machines is up to 40 000 cells per second,
making it feasible to screen libraries containing on the order of 109 variants.
3.4 Selected Industrial Examples
One of the main caveats that come with the integration of enzymatic processes into industrial
settings is that enzymes have evolved through Darwinian evolution for the purpose of survival,
not for the purpose of overproducing valuable pharmaceutical products in harsh industrial
settings. While an enzyme may have the desired catalytic function for a particular processing
step, it most likely will not have sufficient catalytic properties, stability, specificity, or
enantioselectivity when simply exchanged with a developed chemical route of synthesis. In
addition, the production of the enzyme itself may be economically infeasible due to low
expression or solubility. Therefore, there are now many examples of utilizing library creation
and screening tools to enhance enzymes for industrial applications. A small sampling of recent
examples is listed below for the improvement of various enzymatic functions, with further
references for more in-depth study provided for the reader’s benefit.
3.4.1 Activity
The activity of an enzyme is crucial to the productivity of an industrial process. Many
pharmaceutically relevant enzymatic processes utilize enzymes with multiple domains that
catalyze different reactions. One example is a hybrid, nonribosomal peptide synthase–polyketide synthase (NRPS-PKS) from Pantoea agglomerans [64], which produces the broadspectrum antibiotic andrimid [65,66], an acetyl-CoA carboxylase inhibitor. Based on the
proposed pathway for andrimid biosynthesis, the valine-specific A domain of the AdmK
protein was replaced with CytC1, a 2-aminobutyrate-incorporating A domain from the
Streptomyces sp. RK95-74 cytotrienin NRPS-PKS. The CtyC1 domain was chosen due to
its broad specificity profile. While andrimid was still produced with AdmK-CtyC1 hybrid, the
Directed Enzyme Evolution and High-Throughput Screening
57
Figure 3.4 Improvement of the activity of chimeric NRPSs using directed evolution. (1) A heterologous A domain is swapped into an NRPS, typically resulting in a significant loss of synthetase activity.
(2) A library of chimeric synthetase mutants is constructed in which the heterologous A domain has been
diversified (for example, by error-prone PCR). (3) The library is subjected to an in vivo screen for
production of the unnatural nonribosomal peptide derivative. (4) Clones showing improved production
are characterized and subjected to further rounds of diversification and screening
level was reduced by 32-fold compared with the wild-type AdmK. Several rounds of errorprone PCR were performed, and the resulting mutant AdmK-CtyC1 enzyme produced 10.7fold more andrimid than the wild-type AdmK-CtyC1 and only threefold less than the wild-type
AdmK. The screen was also extended to test for the increased production of andrimid
derivatives produced by the new CtyC1 A domain (Figure 3.4). The results indicated that
chimeric NRPSs can be functionally improved, as well as reach activity levels approaching
those of the wild-type counterparts [67].
Cephalosporins are a class of antibiotic produced via the intermediate 7-aminocephalosporanic acid (7-ACA), or 7-aminodesacetoxycephalosporanic acid (7-ADCA). Directed
evolution has been used to improve the activity of cephalosporin acylases to produce these
intermediates from adipyl-7-ACA or cephalosporin C [68]. Using site-directed saturation
mutagenesis and a selection system whereby the E. coli host is dependent on leucine liberated
from derivatives of the cephalosporin side-chains, a mutant was found that increased the
catalytic efficiency toward adipyl-7-ADCA by 36-fold.
Oxidoreductases are a family of enzymes that catalyze a number of industrially important
reactions, but they often require additional nicotinamide (NADH or NADPH) cofactors which
58
Biocatalysis for the Pharmaceutical Industry
are too expensive to supply stoichiometrically. Regeneration of the reduced cofactor form was
accomplished by Woodyer et al. [69] by linking an oxidoreductase reaction to phosphite
dehydrogenase, an enzyme that oxidizes inert phosphite to phosphate with concomitant
reduction of NAD(P) to NAD(P)H. However, the wild-type enzyme had relatively low activity.
Through several rounds of random mutagenesis, the activity of the wild-type enzyme was
improved by sixfold. The advantage of the wild-type enzyme was demonstrated using the
model industrial bioconversion reaction of trimethylpyruvate to L-tert-leucine, an unnatural
amino acid derivative.
3.4.2 Thermostability
An enzyme’s thermostability is generally one of the more difficult characteristics to rationally
improve. Beneficial mutations that lead to increased thermostability can be involved in many
different mechanisms, including solvent interactions, structural support, and electrostatic
balance. The majority of attempts to enhance the thermostability of an enzyme have, in some
way or another, incorporated random mutagenesis techniques to generate improved enzymes.
However, once improved mutants have been screened for and isolated, combining the different
mutations together can be labor intensive. Through biased mutation-assembling, which
simplified and reduced the workload of incorporating multiple beneficial mutations, Hamamatsu
et al. [20] were able to improve the thermostability of a prolyl endopeptidase from Flavobacterium meningosepticum, resulting in a 1200-fold improvement of the enzyme’s half-life at 60 C.
Another example involved a combination of computational data, rational design, and directed
evolution. Iterative saturation mutagenesis [70] was used on a model enzyme lipase from Bacillus
subtilis to improve its thermostability. Appropriate amino acid residues were chosen based on
X-ray data, in particular the B factors (or B values). These factors reflect smearing of atomic
electron densities with respect to their equilibrium positions as a result of thermal motion and
positional disorder. Amino acids which display large B factors correspond to those which have
pronounced flexibility and are targets for stabilization. Each position was subjected to saturation
mutagenesis, to generate as many single position libraries as were suitable for the experiment.
After screening, the improved mutants found were ranked by position on their contribution to the
enzyme’s improved thermostability. Consecutive rounds of saturation mutagenesis were then
performed, first starting with the position of highest improvement, followed by the second
highest, and so on until all positions had been explored. The B. subtilis lipase was evolved through
this method to two mutants (five and seven total mutations each) that had half-lives of more than
15 h at 55 C, compared with 2 min for the wild-type enzyme. The principle of additive
mutational effects played an important role in this design. Also, the selection of residues was
limited to having crystal structures available, although the same process can be carried out with
template mutants found by other random mutagenesis techniques.
3.4.3 Substrate Specificity
The cholesterol-lowering drug atorvastatin, marketed as Lipitor, is an example where biocatalysis research has been applied extensively and is in industrial use. The enzyme 2-deoxyribose5-phosphate aldolase (DERA) has been a target of directed evolution for the production of
atorvastatin intermediates [8,9,71]. DeSantis and coworkers [8,9] used structure-based
Directed Enzyme Evolution and High-Throughput Screening
59
mutagenesis in an attempt to expand the substrate specificity of the native enzyme. The enzyme
variants were fully purified and assayed in vitro by monitoring the decrease of NADH at 340 nm
using a spectrophotometer. One variant, S238D, showed new activity towards 3-azidopropinaldehyde to form an azido pyranose which is an intermediate in atorvastatin synthesis.
Jennewein et al. [71] also studied this wild-type enzyme, creating a library with error-prone
PCR and recombining positive mutants. By screening mutants with a microplate reader or with
gas chromatography, they managed to increase the synthesis of the intermediate (3R,5S)-6chloro-2,4,6-trideoxyhexapyranoside by 10-fold.
3.4.4 Product Specificity
Polyketides are a structurally diverse group of compounds and one of the richest sources of
pharmaceuticals. The biosynthesis of the polyketide avermectin-analog doramectin (commercially sold by Pfizer as Dectomax) in a mutant Streptomyces avermitilis strain was hampered
by the product specificity of one of the key intermediate enzymes, aveC, in the doramectin
production pathway. Although the particular function of aveC was unknown, the occurrence of
undesirable product CHC-B2 was found to be a result of its dual-product specificity. In an effort
to improve the production ratio of doramectin:CHC-B2, site-specific and error-prone mutagenesis libraries were screened and resulted in several unrelated beneficial mutations that
improved the production ratio up to fourfold [72]. However, much of the screened library had
mutations that adversely affected the production ratio of doramectin. Continuing the efforts to
improve aveC for doramectin production, a new approach was established to curb the complex
and time-consuming initial screens, due in large part to the long growth periods of the
S. avermitilis host strain and involved the electrospray ionization tandem mass spectrometry
method to detect the resultant products. A semi-synthetic shuffling method [73] was utilized to
allow for screening of the diversity of mutations generated in the initial rounds while reducing
the library size to be screened. After three rounds of semi-synthetic shuffling, an improved
mutant containing 10 mutations was inserted into production strains, and doramectin ratios
were found to be improved more than 23-fold over the wild type [74].
3.4.5 Enantioselectivity
The chiral molecule (R)-4-cyano-3-hydroxybutyric acid is another intermediate in the synthesis of atorvastatin (Figure 3.5), and its production enzymatically has been targeted by a
number of groups [8,9,75]. DeSantis and coworkers [8,9] used a route involving the hydrolysis
of 3-hydroxyglutaryl nitrile by a nitrilase enzyme. GSSM was used to increase the enantioselectivity under high substrate conditions. A chiral substrate was synthesized whereby one
nitrile group contained the nitrogen isotope 15 N, meaning that the R- and S-products differed by
one mass unit, which was detected by mass spectrometry. Screening of over 30 000 variants
indicated the best mutant contained the Ala190His mutation, which increased the enantiomeric
excess (ee) to 98.1% from 87.8% and had complete conversion in 15 h as opposed to 24 h when
2.25 M hydroxyglutaryl nitrile was used. Fox et al. [75] used a halohydrin dehydrogenase to
produce ethyl (R)-4-cyano-3-hydroxybutyrate, a starting point for atorvastatin synthesis. The
enzyme from Agrobacterium radiobacter acts on ethyl (S)-4-chloro-3-hydroxybutyrate,
forming the product but initially at a low rate. A statistical model based on protein
structure–activity relationships was incorporated, and 18 rounds of screening were carried
60
Biocatalysis for the Pharmaceutical Industry
Figure 3.5 Directed evolution applied to enzymes for the production of atorvastatin synthesis intermediates. The substrates and products of the evolved enzymes are shown
out. The final mutant showed a 4000-fold improvement in volumetric production, with the ethyl
(R)-4-cyano-3-hydroxybutyrate product being 99.5% pure and with an ee of over 99.9%.
Epothilones are a class of molecules that show anticancer activity. Production of a synthetic
intermediate was investigated through the action of an esterase on various sterically hindered
3-hydroxy esters [76]. No initial activity was observed, so a Pseudomonas fluorescens esterase
was transformed into a mutator strain Epicurian coli and screened using an indicator in the
growth plates that would produce a red color if hydrolysis occurred. An ee of 25% was achieved
from a variant containing two mutations.
3.5 Conclusions and Future Directions
The complexity of today’s pharmaceutical compounds and an increasing awareness of the
environmental impact of traditional chemical syntheses have opened the door to biocatalysis.
Directed evolution is an integral tool in the development of synthetic enzymes, ensuring they
are suitable for use in an industrial setting. The past success of this approach indicates that it
will continue to provide many examples of safe and efficient production of chemical
intermediates and medical compounds.
Directed Enzyme Evolution and High-Throughput Screening
61
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