Protein engineering is the process of developing useful or valuable proteins through the design and production of unnatural polypeptides, often by altering amino acid sequences found in nature. It is a young discipline, with much research taking place into the understanding of protein folding and recognition for protein design principles. It has been used to improve the function of many enzymes for industrial catalysis. It is also a product and services market, with an estimated value of $168 billion by 2017.
There are two general strategies for protein engineering: rational protein design and directed evolution. These methods are not mutually exclusive; researchers will often apply both. In the future, more detailed knowledge of protein structure and function, and advances in high-throughput screening,
may greatly expand the abilities of protein engineering. Eventually,
even unnatural amino acids may be included, via newer methods, such as expanded genetic code, that allow encoding novel amino acids in genetic code.
The applications in numerous fields, including medicine and industrial bioprocessing, are vast and numerous.
Approaches
Rational design
In rational protein design, a scientist uses detailed knowledge of
the structure and function of a protein to make desired changes. In
general, this has the advantage of being inexpensive and technically
easy, since site-directed mutagenesis
methods are well-developed. However, its major drawback is that
detailed structural knowledge of a protein is often unavailable, and,
even when available, it can be very difficult to predict the effects of
various mutations since structural information most often provide a
static picture of a protein structure. However, programs such as Folding@home and Foldit have utilized crowdsourcing techniques in order to gain insight into the folding motifs of proteins.
Computational protein design algorithms
seek to identify novel amino acid sequences that are low in energy when
folded to the pre-specified target structure. While the
sequence-conformation space that needs to be searched is large, the most
challenging requirement for computational protein design is a fast, yet
accurate, energy function that can distinguish optimal sequences from
similar suboptimal ones.
Multiple sequence alignment
Without structural information about a protein, sequence analysis is
often useful in elucidating information about the protein. These
techniques involve alignment of target protein sequences with other
related protein sequences. This alignment can show which amino acids are
conserved between species and are important for the function of the
protein. These analyses can help to identify hot spot amino acids that
can serve as the target sites for mutations. Multiple sequence alignment
utilizes data bases such as PREFAB, SABMARK, OXBENCH, IRMBASE, and
BALIBASE in order to cross reference target protein sequences with known
sequences. Multiple sequence alignment techniques are listed below.
This method begins by performing pair wise alignment of sequences using k-tuple or Needleman–Wunsch
methods. These methods calculate a matrix that depicts the pair wise
similarity among the sequence pairs. Similarity scores are then
transformed into distance scores that are used to produce a guide tree
using the neighbor joining method. This guide tree is then employed to
yield a multiple sequence alignment.
Clustal omega
This method is capable of aligning up to 190,000 sequences by
utilizing the k-tuple method. Next sequences are clustered using the
mBed and k-means methods. A guide tree is then constructed using the UPGMA method that is used by the HH align package. This guide tree is used to generate multiple sequence alignments.
MAFFT
This method utilizes fast Fourier transform (FFT) that converts amino
acid sequences into a sequence composed of volume and polarity values
for each amino acid residue. This new sequence is used to find
homologous regions.
K-Align
This method utilizes the Wu-Manber approximate string matching algorithm to generate multiple sequence alignments.
Multiple sequence comparison by log expectation (MUSCLE)
This method utilizes Kmer and Kimura distances to generate multiple sequence alignments.
T-Coffee
This method utilizes tree based consistency objective functions for
alignment evolution. This method has been shown to be 5–10% more
accurate than Clustal W.
Coevolutionary analysis
Coevolutionary analysis is also known as correlated mutation,
covariation, or co-substitution. This type of rational design involves
reciprocal evolutionary changes at evolutionarily interacting loci.
Generally this method begins with the generation of a curated multiple
sequence alignments for the target sequence. This alignment is then
subjected to manual refinement that involves removal of highly gapped
sequences, as well as sequences with low sequence identity. This step
increases the quality of the alignment. Next, the manually processed
alignment is utilized for further coevolutionary measurements using
distinct correlated mutation algorithms. These algorithms result in a
coevolution scoring matrix. This matrix is filtered by applying various
significance tests to extract significant coevolution values and wipe
out background noise. Coevolutionary measurements are further evaluated
to assess their performance and stringency. Finally, the results from
this coevolutionary analysis are validated experimentally.
Structural prediction
De novo generation of protein benefits from knowledge of
existing protein structures. This knowledge of existing protein
structure assists with the prediction of new protein structures. Methods
for protein structure prediction fall under one of the four following
classes: ab initio, fragment based methods, homology modeling, and protein threading.
Ab initio
These methods involve free modeling without using any structural information about the template. Ab initio
methods are aimed at prediction of the native structures of proteins
corresponding to the global minimum of its free energy. some examples of
ab initio methods are AMBER, GROMOS, GROMACS, CHARMM, OPLS, and ENCEPP12. General steps for ab initio
methods begin with the geometric representation of the protein of
interest. Next, a potential energy function model for the protein is
developed. This model can be created using either molecular mechanics
potentials or protein structure derived potential functions. Following
the development of a potential model, energy search techniques including
molecular dynamic simulations, Monte Carlo simulations and genetic
algorithms are applied to the protein.
Fragment based
These methods use database information regarding structures to match
homologous structures to the created protein sequences. These homologous
structures are assembled to give compact structures using scoring and
optimization procedures, with the goal of achieving the lowest potential
energy score. Webservers for fragment information are I-TASSER,
ROSETTA, ROSETTA @ home, FRAGFOLD, CABS fold, PROFESY, CREF, QUARK,
UNDERTAKER, HMM, and ANGLOR.
Homology modeling
These methods are based upon the homology of proteins. These methods
are also known as comparative modeling. The first step in homology
modeling is generally the identification of template sequences of known
structure which are homologous to the query sequence. Next the query
sequence is aligned to the template sequence. Following the alignment,
the structurally conserved regions are modeled using the template
structure. This is followed by the modeling of side chains and loops
that are distinct from the template. Finally the modeled structure
undergoes refinement and assessment of quality. Servers that are
available for homology modeling data are listed here: SWISS MODEL,
MODELLER, ReformAlign, PyMOD, TIP-STRUCTFAST, COMPASS, 3d-PSSM, SAMT02,
SAMT99, HHPRED, FAGUE, 3D-JIGSAW, META-PP, ROSETTA, and I-TASSER.
Protein threading
Protein threading can be used when a reliable homologue for the query
sequence cannot be found. This method begins by obtaining a query
sequence and a library of template structures. Next, the query sequence
is threaded over known template structures. These candidate models are
scored using scoring functions. These are scored based upon potential
energy models of both query and template sequence. The match with the
lowest potential energy model is then selected. Methods and servers for
retrieving threading data and performing calculations are listed here:
GenTHREADER, pGenTHREADER, pDomTHREADER, ORFEUS, PROSPECT,
BioShell-Threading, FFASO3, RaptorX, HHPred, LOOPP server, Sparks-X,
SEGMER, THREADER2, ESYPRED3D, LIBRA, TOPITS, RAPTOR, COTH, MUSTER.
For more information on rational design see site-directed mutagenesis.
Multivalent binding
Multivalent binding can be used to increase the binding specificity and affinity through avidity
effects. Having multiple binding domains in a single biomolecule or
complex increases the likelihood of other interactions to occur via
individual binding events. Avidity or effective affinity can be much
higher than the sum of the individual affinities providing a cost and
time-effective tool for targeted binding.
Multivalent proteins
Multivalent proteins are relatively easy to produce by
post-translational modifications or multiplying the protein-coding DNA
sequence. The main advantage of multivalent and multispecific proteins
is that they can increase the effective affinity for a target of a known
protein. In the case of an inhomogeneous target using a combination of
proteins resulting in multispecific binding can increase specificity,
which has high applicability in protein therapeutics.
The most common example for multivalent binding are the
antibodies, and there is extensive research for bispecific antibodies.
Applications of bispecific antibodies cover a broad spectrum that
includes diagnosis, imaging, prophylaxis, and therapy.
Directed evolution
In directed evolution, random mutagenesis, e.g. by error-prone PCR or sequence saturation mutagenesis,
is applied to a protein, and a selection regime is used to select
variants having desired traits. Further rounds of mutation and
selection are then applied. This method mimics natural evolution and, in general, produces superior results to rational design. An added process, termed DNA shuffling, mixes and matches pieces of successful variants to produce better results. Such processes mimic the recombination that occurs naturally during sexual reproduction.
Advantages of directed evolution are that it requires no prior
structural knowledge of a protein, nor is it necessary to be able to
predict what effect a given mutation will have. Indeed, the results of
directed evolution experiments are often surprising in that desired
changes are often caused by mutations that were not expected to have
some effect. The drawback is that they require high-throughput screening, which is not feasible for all proteins. Large amounts of recombinant DNA
must be mutated and the products screened for desired traits. The
large number of variants often requires expensive robotic equipment to
automate the process. Further, not all desired activities can be
screened for easily.
Natural Darwinian evolution can be effectively imitated in the
lab toward tailoring protein properties for diverse applications,
including catalysis. Many experimental technologies exist to produce
large and diverse protein libraries and for screening or selecting
folded, functional variants. Folded proteins arise surprisingly
frequently in random sequence space, an occurrence exploitable in
evolving selective binders and catalysts. While more conservative than
direct selection from deep sequence space, redesign of existing proteins
by random mutagenesis and selection/screening is a particularly robust
method for optimizing or altering extant properties. It also represents
an excellent starting point for achieving more ambitious engineering
goals. Allying experimental evolution with modern computational methods
is likely the broadest, most fruitful strategy for generating functional
macromolecules unknown to nature.
The main challenges of designing high quality mutant libraries
have shown significant progress in the recent past. This progress has
been in the form of better descriptions of the effects of mutational
loads on protein traits. Also computational approaches have shown large
advances in the innumerably large sequence space to more manageable
screenable sizes, thus creating smart libraries of mutants. Library size
has also been reduced to more screenable sizes by the identification of
key beneficial residues using algorithms for systematic recombination.
Finally a significant step forward toward efficient reengineering of
enzymes has been made with the development of more accurate statistical
models and algorithms quantifying and predicting coupled mutational
effects on protein functions.
Generally, directed evolution may be summarized as an iterative
two step process which involves generation of protein mutant libraries,
and high throughput screening processes to select for variants with
improved traits. This technique does not require prior knowledge of the
protein structure and function relationship. Directed evolution utilizes
random or focused mutagenesis to generate libraries of mutant proteins.
Random mutations can be introduced using either error prone PCR, or
site saturation mutagenesis. Mutants may also be generated using
recombination of multiple homologous genes. Nature has evolved a limited
number of beneficial sequences. Directed evolution makes it possible to
identify undiscovered protein sequences which have novel functions.
This ability is contingent on the proteins ability to tolerant amino
acid residue substitutions without compromising folding or stability.
Directed evolution methods can be broadly categorized into two strategies, asexual and sexual methods.
Asexual methods
Asexual methods do not generate any cross links between parental
genes. Single genes are used to create mutant libraries using various
mutagenic techniques. These asexual methods can produce either random or
focused mutagenesis.
Random mutagenesis
Random mutagenic methods produce mutations at random throughout the
gene of interest. Random mutagenesis can introduce the following types
of mutations: transitions, transversions, insertions, deletions,
inversion, missense, and nonsense. Examples of methods for producing
random mutagenesis are below.
Error prone PCR
Error prone PCR utilizes the fact that Taq DNA polymerase lacks 3' to
5' exonuclease activity. This results in an error rate of 0.001–0.002%
per nucleotide per replication. This method begins with choosing the
gene, or the area within a gene, one wishes to mutate. Next, the extent
of error required is calculated based upon the type and extent of
activity one wishes to generate. This extent of error determines the
error prone PCR strategy to be employed. Following PCR, the genes are
cloned into a plasmid and introduced to competent cell systems. These
cells are then screened for desired traits. Plasmids are then isolated
for colonies which show improved traits, and are then used as templates
the next round of mutagenesis. Error prone PCR shows biases for certain
mutations relative to others. Such as biases for transitions over
transversions.
Rates of error in PCR can be increased in the following ways:
- Increase concentration of magnesium chloride, which stabilizes non complementary base pairing.
- Add manganese chloride to reduce base pair specificity.
- Increased and unbalanced addition of dNTPs.
- Addition of base analogs like dITP, 8 oxo-dGTP, and dPTP.
- Increase concentration of Taq polymerase.
- Increase extension time.
- Increase cycle time.
- Use less accurate Taq polymerase.
Also see polymerase chain reaction for more information.
Rolling circle error-prone PCR
This PCR method is based upon rolling circle amplification, which is
modeled from the method that bacteria use to amplify circular DNA. This
method results in linear DNA duplexes. These fragments contain tandem
repeats of circular DNA called concatamers, which can be transformed
into bacterial strains. Mutations are introduced by first cloning the
target sequence into an appropriate plasmid. Next, the amplification
process begins using random hexamer primers and Φ29 DNA polymerase
under error prone rolling circle amplification conditions. Additional
conditions to produce error prone rolling circle amplification are 1.5
pM of template DNA, 1.5 mM MnCl2 and a 24 hour reaction time. MnCl2
is added into the reaction mixture to promote random point mutations in
the DNA strands. Mutation rates can be increased by increasing the
concentration of MnCl2, or by decreasing concentration of the
template DNA. Error prone rolling circle amplification is advantageous
relative to error prone PCR because of its use of universal random
hexamer primers, rather than specific primers. Also the reaction
products of this amplification do not need to be treated with ligases or
endonucleases. This reaction is isothermal.
Chemical mutagenesis
Chemical mutagenesis involves the use of chemical agents to introduce
mutations into genetic sequences. Examples of chemical mutagens follow.
Sodium bisulfate is effective at mutating G/C rich genomic
sequences. This is because sodium bisulfate catalyses deamination of
unmethylated cytosine to uracil.
Ethyl methane sulfonate alkylates guanidine residues. This alteration causes errors during DNA replication.
Nitrous acid causes transversion by de-amination of adenine and cytosine.
The dual approach to random chemical mutagenesis is an iterative two step process. First it involves the in vivo
chemical mutagenesis of the gene of interest via EMS. Next, the treated
gene is isolated and cloning into an untreated expression vector in
order to prevent mutations in the plasmid backbone. This technique preserves the plasmids genetic properties.
Targeting glycosylases to embedded arrays for mutagenesis (TaGTEAM)
This method has been used to create targeted in vivo
mutagenesis in yeast. This method involves the fusion of a
3-methyladenine DNA glycosylase to tetR DNA-binding domain. This has
been shown to increase mutation rates by over 800 time in regions of the
genome containing tetO sites.
Mutagenesis by random insertion and deletion
This method involves alteration in length of the sequence via
simultaneous deletion and insertion of chunks of bases of arbitrary
length. This method has been shown to produce proteins with new
functionalities via introduction of new restriction sites, specific
codons, four base codons for non-natural amino acids.
Transposon based random mutagenesis
Recently many methods for transposon based random mutagenesis have
been reported. This methods include, but are not limited to the
following: PERMUTE-random circular permutation, random protein
truncation, random nucleotide triplet substitution, random
domain/tag/multiple amino acid insertion, codon scanning mutagenesis,
and multicodon scanning mutagenesis. These aforementioned techniques all
require the design of mini-Mu transposons. Thermo scientific
manufactures kits for the design of these transposons.
Random mutagenesis methods altering the target DNA length
These methods involve altering gene length via insertion and deletion
mutations. An example is the tandem repeat insertion (TRINS) method.
This technique results in the generation of tandem repeats of random
fragments of the target gene via rolling circle amplification and
concurrent incorporation of these repeats into the target gene.
Mutator strains
Mutator strains are bacterial cell lines which are deficient in one
or more DNA repair mechanisms. An example of a mutator strand is the E.
coli XL1-RED. This subordinate strain of E. coli is deficient in the MutS, MutD, MutT
DNA repair pathways. Use of mutator strains is useful at introducing
many types of mutation; however, these strains show progressive sickness
of culture because of the accumulation of mutations in the strains own
genome.
Focused mutagenesis
Focused mutagenic methods produce mutations at predetermined amino
acid residues. These techniques require and understanding of the
sequence-function relationship for the protein of interest.
Understanding of this relationship allows for the identification of
residues which are important in stability, stereoselectivity, and
catalytic efficiency. Examples of methods that produce focused mutagenesis are below.
Site saturation mutagenesis
Site saturation mutagenesis is a PCR based method used to target
amino acids with significant roles in protein function. The two most
common techniques for performing this are whole plasmid single PCR, and
overlap extension PCR.
Whole plasmid single PCR is also referred to as site directed
mutagenesis (SDM). SDM products are subjected to Dpn endonuclease
digestion. This digestion results in cleavage of only the parental
strand, because the parental strand contains a GmATC which is methylated
at N6 of adenine. SDM does not work well for large plasmids of over ten
kilobases. Also, this method is only capable of replacing two
nucleotides at a time.
Overlap extension PCR requires the use of two pairs of primers.
One primer in each set contains a mutation. A first round of PCR using
these primer sets is performed and two double stranded DNA duplexes are
formed. A second round of PCR is then performed in which these duplexes
are denatured and annealed with the primer sets again to produce
heteroduplexes, in which each strand has a mutation. Any gaps in these
newly formed heteroduplexes are filled with DNA polymerases and further
amplified.
Sequence saturation mutagenesis (SeSaM)
Sequence saturation mutagenesis
results in the randomization of the target sequence at every nucleotide
position. This method begins with the generation of variable length DNA
fragments tailed with universal bases via the use of template
transferases at the 3' termini. Next, these fragments are extended to
full length using a single stranded template. The universal bases are
replaced with a random standard base, causing mutations. There are
several modified versions of this method such as SeSAM-Tv-II, SeSAM-Tv+,
and SeSAM-III.
Single primer reactions in parallel (SPRINP)
This site saturation mutagenesis method involves two separate PCR
reaction. The first of which uses only forward primers, while the second
reaction uses only reverse primers. This avoids the formation of primer
dimer formation.
Mega primed and ligase free focused mutagenesis
This site saturation mutagenic technique begins with one mutagenic
oligonucleotide and one universal flanking primer. These two reactants
are used for an initial PCR cycle. Products from this first PCR cycle
are used as mega primers for the next PCR.
Ω-PCR
This site saturation mutagenic method is based on overlap extension
PCR. It is used to introduce mutations at any site in a circular
plasmid.
PFunkel-ominchange-OSCARR
This method utilizes user defined site directed mutagenesis at single
or multiple sites simultaneously. OSCARR is an acronym for one pot simple methodology for cassette randomization and recombination.
This randomization and recombination results in randomization of
desired fragments of a protein. Omnichange is a sequence independent,
multisite saturation mutagenesis which can saturate up to five
independent codons on a gene.
Trimer-dimer mutagenesis
This method removes redundant codons and stop codons.
Cassette mutagenesis
This is a PCR based method. Cassette mutagenesis
begins with the synthesis of a DNA cassette containing the gene of
interest, which is flanked on either side by restriction sites. The
endonuclease which cleaves these restriction sites also cleaves sites in
the target plasmid. The DNA cassette and the target plasmid are both
treated with endonucleases to cleave these restriction sites and create
sticky ends. Next the products from this cleavage are ligated together,
resulting in the insertion of the gene into the target plasmid. An
alternative form of cassette mutagenesis called combinatorial cassette
mutagenesis is used to identify the functions of individual amino acid
residues in the protein of interest. Recursive ensemble mutagenesis then
utilizes information from previous combinatorial cassette mutagenesis.
Codon cassette mutagenesis allows you to insert or replace a single
codon at a particular site in double stranded DNA.
Sexual methods
Sexual methods of directed evolution involve in vitro recombination which mimic natural in vivo recombination. Generally these techniques require high sequence homology
between parental sequences. These techniques are often used to
recombine two different parental genes, and these methods do create
cross overs between these genes.
In vitro homologous recombination
Homologous recombination can be categorized as either in vivo or in vitro. In vitro homologous recombination mimics natural in vivo recombination. These in vitro
recombination methods require high sequence homology between parental
sequences. These techniques exploit the natural diversity in parental
genes by recombining them to yield chimeric genes. The resulting chimera
show a blend of parental characteristics.
DNA shuffling
This in vitro technique was one of the first techniques in the
era of recombination. It begins with the digestion of homologous
parental genes into small fragments by DNase1. These small fragments are
then purified from undigested parental genes. Purified fragments are
then reassembled using primer-less PCR. This PCR involves homologous
fragments from different parental genes priming for each other,
resulting in chimeric DNA. The chimeric DNA of parental size is then
amplified using end terminal primers in regular PCR.
Random priming in vitro recombination (RPR)
This in vitro homologous recombination method begins with the
synthesis of many short gene fragments exhibiting point mutations using
random sequence primers. These fragments are reassembled to full length
parental genes using primer-less PCR. These reassembled sequences are
then amplified using PCR and subjected to further selection processes.
This method is advantageous relative to DNA shuffling because there is
no use of DNase1, thus there is no bias for recombination next to a
pyrimidine nucleotide. This method is also advantageous due to its use
of synthetic random primers which are uniform in length, and lack
biases. Finally this method is independent of the length of DNA template
sequence, and requires a small amount of parental DNA.
This method generates chimeric genes directly from metagenomic
samples. It begins with isolation of the desired gene by functional
screening from metagenomic DNA sample. Next, specific primers are
designed and used to amplify the homologous genes from different
environmental samples. Finally, chimeric libraries are generated to
retrieve the desired functional clones by shuffling these amplified
homologous genes.
Staggered extension process (StEP)
This in vitro method is based on template switching to
generate chimeric genes. This PCR based method begins with an initial
denaturation of the template, followed by annealing of primers and a
short extension time. All subsequent cycle generate annealing between
the short fragments generated in previous cycles and different parts of
the template. These short fragments and the templates anneal together
based on sequence complementarity. This process of fragments annealing
template DNA is known as template switching. These annealed fragments
will then serve as primers for further extension. This method is carried
out until the parental length chimeric gene sequence is obtained.
Execution of this method only requires flanking primers to begin. There
is also no need for Dnase1 enzyme.
Random chimeragenesis on transient templates (RACHITT)
This method has been shown to generate chimeric gene libraries with
an average of 14 crossovers per chimeric gene. It begins by aligning
fragments from a parental top strand onto the bottom strand of a uracil
containing template from a homologous gene. 5' and 3' overhang flaps are
cleaved and gaps are filled by the exonuclease and endonuclease
activities of Pfu and taq DNA polymerases. The uracil containing
template is then removed from the heteroduplex by treatment with a
uracil DNA glcosylase, followed by further amplification using PCR. This
method is advantageous because it generates chimeras with relatively
high crossover frequency. However it is somewhat limited due to the
complexity and the need for generation of single stranded DNA and uracil
containing single stranded template DNA.
Synthetic shuffling
Shuffling of synthetic degenerate oligonucleotides adds flexibility
to shuffling methods, since oligonucleotides containing optimal codons
and beneficial mutations can be included.
In vivo Homologous Recombination
Cloning performed in yeast involves PCR dependent reassembly of
fragmented expression vectors. These reassembled vectors are then
introduced to, and cloned in yeast. Using yeast to clone the vector
avoids toxicity and counter-selection that would be introduced by
ligation and propagation in E. coli.
Mutagenic organized recombination process by homologous in vivo grouping (MORPHING)
This method introduces mutations into specific regions of genes while
leaving other parts intact by utilizing the high frequency of
homologous recombination in yeast.
Phage-assisted continuous evolution (PACE)
This method utilizes a bacteriophage with a modified life cycle to
transfer evolving genes from host to host. The phage's life cycle is
designed in such a way that the transfer is correlated with the activity
of interest from the enzyme. This method is advantageous because it
requires minimal human intervention for the continuous evolution of the
gene.
In vitro non-homologous recombination methods
These methods are based upon the fact that proteins can exhibit similar structural identity while lacking sequence homology.
Exon shuffling
Exon shuffling is the combination of exons from different proteins by
recombination events occurring at introns. Orthologous exon shuffling
involves combining exons from orthologous genes from different species.
Orthologous domain shuffling involves shuffling of entire protein
domains from orthologous genes from different species. Paralogous exon
shuffling involves shuffling of exon from different genes from the same
species. Paralogous domain shuffling involves shuffling of entire
protein domains from paralogous proteins from the same species.
Functional homolog shuffling involves shuffling of non-homologous
domains which are functional related. All of these processes being with
amplification of the desired exons from different genes using chimeric
synthetic oligonucleotides. This amplification products are then
reassembled into full length genes using primer-less PCR. During these
PCR cycles the fragments act as templates and primers. This results in
chimeric full length genes, which are then subjected to screening.
Incremental truncation for the creation of hybrid enzymes (ITCHY)
Fragments of parental genes are created using controlled digestion by
exonuclease III. These fragments are blunted using endonuclease, and
are ligated to produce hybrid genes. THIOITCHY is a modified ITCHY
technique which utilized nucleotide triphosphate analogs such as α-phosphothioate dNTPs.
Incorporation of these nucleotides blocks digestion by exonuclease
III. This inhibition of digestion by exonuclease III is called spiking.
Spiking can be accomplished by first truncating genes with exonuclease
to create fragments with short single stranded overhangs. These
fragments then serve as templates for amplification by DNA polymerase in
the presence of small amounts of phosphothioate dNTPs. These resulting
fragments are then ligated together to form full length genes.
Alternatively the intact parental genes can be amplified by PCR in the
presence of normal dNTPs and phosphothioate dNTPs. These full length
amplification products are then subjected to digestion by an
exonuclease. Digestion will continue until the exonuclease encounters an
α-pdNTP, resulting in fragments of different length. These fragments
are then ligated together to generate chimeric genes.
SCRATCHY
This method generates libraries of hybrid genes inhibiting multiple
crossovers by combining DNA shuffling and ITCHY. This method begins with
the construction of two independent ITCHY libraries. The first with
gene A on the N-terminus. And the other having gene B on the N-terminus.
These hybrid gene fragments are separated using either restriction
enzyme digestion or PCR with terminus primers via agarose gel
electrophoresis. These isolated fragments are then mixed together and
further digested using DNase1. Digested fragments are then reassembled
by primerless PCR with template switching.
Recombined extension on truncated templates (RETT)
This method generates libraries of hybrid genes by template switching
of uni-directionally growing polynucleotides in the presence of single
stranded DNA fragments as templates for chimeras. This method begins
with the preparation of single stranded DNA fragments by reverse
transcription from target mRNA. Gene specific primers are then annealed
to the single stranded DNA. These genes are then extended during a PCR
cycle. This cycle is followed by template switching and annealing of the
short fragments obtained from the earlier primer extension to other
single stranded DNA fragments. This process is repeated until full
length single stranded DNA is obtained.
Sequence homology-independent protein recombination (SHIPREC)
This method generates recombination between genes with little to no
sequence homology. These chimeras are fused via a linker sequence
containing several restriction sites. This construct is then digested
using DNase1. Fragments are made are made blunt ended using S1 nuclease.
These blunt end fragments are put together into a circular sequence by
ligation. This circular construct is then linearized using restriction
enzymes for which the restriction sites are present in the linker
region. This results in a library of chimeric genes in which
contribution of genes to 5' and 3' end will be reversed as compared to
the starting construct.
Sequence independent site directed chimeragenesis (SISDC)
This method results in a library of genes with multiple crossovers
from several parental genes. This method does not require sequence
identity among the parental genes. This does require one or two
conserved amino acids at every crossover position. It begins with
alignment of parental sequences and identification of consensus regions
which serve as crossover sites. This is followed by the incorporation of
specific tags containing restriction sites followed by the removal of
the tags by digestion with Bac1, resulting in genes with cohesive ends.
These gene fragments are mixed and ligated in an appropriate order to
form chimeric libraries.
Degenerate homo-duplex recombination (DHR)
This method begins with alignment of homologous genes, followed by
identification of regions of polymorphism. Next the top strand of the
gene is divided into small degenerate oligonucleotides. The bottom
strand is also digested into oligonucleotides to serve as scaffolds.
These fragments are combined in solution are top strand oligonucleotides
are assembled onto bottom strand oligonucleotides. Gaps between these
fragments are filled with polymerase and ligated.
Random multi-recombinant PCR (RM-PCR)
This method involves the shuffling of plural DNA fragments without
homology, in a single PCR. This results in the reconstruction of
complete proteins by assembly of modules encoding different structural
units.
User friendly DNA recombination (USERec)
This method begins with the amplification of gene fragments which
need to be recombined, using uracil dNTPs. This amplification solution
also contains primers, PfuTurbo, and Cx Hotstart DNA polymerase.
Amplified products are next incubated with USER enzyme. This enzyme
catalyzes the removal of uracil residues from DNA creating single base
pair gaps. The USER enzyme treated fragments are mixed and ligated using
T4 DNA ligase and subjected to Dpn1 digestion to remove the template
DNA. These resulting dingle stranded fragments are subjected to
amplification using PCR, and are transformed into E. coli.
Golden Gate shuffling (GGS) recombination
This method allows you to recombine at least 9 different fragments in
an acceptor vector by using type 2 restriction enzyme which cuts
outside of the restriction sites. It begins with sub cloning of
fragments in separate vectors to create Bsa1 flanking sequences on both
sides. These vectors are then cleaved using type II restriction enzyme
Bsa1, which generates four nucleotide single strand overhangs. Fragments
with complementary overhangs are hybridized and ligated using T4 DNA
ligase. Finally these constructs are then transformed into E. coli
cells, which are screened for expression levels.
Phosphoro thioate-based DNA recombination method (PRTec)
This method can be used to recombine structural elements or entire
protein domains. This method is based on phosphorothioate chemistry
which allows the specific cleavage of phosphorothiodiester bonds. The
first step in the process begins with amplification of fragments that
need to be recombined along with the vector backbone. This amplification
is accomplished using primers with phosphorothiolated nucleotides at 5'
ends. Amplified PCR products are cleaved in an ethanol-iodine solution
at high temperatures. Next these fragments are hybridized at room
temperature and transformed into E. coli which repair any nicks.
Integron
This system is based upon a natural site specific recombination
system in E. coli. This system is called the integron system, and
produces natural gene shuffling. This method was used to construct and
optimize a functional tryptophan biosynthetic operon in trp-deficient E.
coli by delivering individual recombination cassettes or trpA-E genes
along with regulatory elements with the integron system.
Y-Ligation based shuffling (YLBS)
This method generates single stranded DNA strands, which encompass a
single block sequence either at the 5' or 3' end, complementary
sequences in a stem loop region, and a D branch region serving as a
primer binding site for PCR. Equivalent amounts of both 5' and 3' half
strands are mixed and formed a hybrid due to the complementarity in the
stem region. Hybrids with free phosphorylated 5' end in 3' half strands
are then ligated with free 3' ends in 5' half strands using T4 DNA
ligase in the presence of 0.1 mM ATP. Ligated products are then
amplified by two types of PCR to generate pre 5' half and pre 3' half
PCR products. These PCR product are converted to single strands via
avidin-biotin binding to the 5' end of the primes containing stem
sequences that were biotin labeled. Next, biotinylated 5' half strands
and non-biotinylated 3' half strands are used as 5' and 3' half strands
for the next Y-ligation cycle.
Semi-rational design
Semi-rational design uses information about a proteins sequence,
structure and function, in tandem with predictive algorithms. Together
these are used to identify target amino acid residues which are most
likely to influence protein function. Mutations of these key amino acid
residues create libraries of mutant proteins that are more likely to
have enhanced properties.
Advances in semi-rational enzyme engineering and de novo enzyme
design provide researchers with powerful and effective new strategies to
manipulate biocatalysts. Integration of sequence and structure based
approaches in library design has proven to be a great guide for enzyme
redesign. Generally, current computational de novo and redesign methods
do not compare to evolved variants in catalytic performance. Although
experimental optimization may be produced using directed evolution,
further improvements in the accuracy of structure predictions and
greater catalytic ability will be achieved with improvements in design
algorithms. Further functional enhancements may be included in future
simulations by integrating protein dynamics.
Biochemical and biophysical studies, along with fine-tuning of
predictive frameworks will be useful to experimentally evaluate the
functional significance of individual design features. Better
understanding of these functional contributions will then give feedback
for the improvement of future designs.
Directed evolution will likely not be replaced as the method of
choice for protein engineering, although computational protein design
has fundamentally changed the way protein engineering can manipulate
bio-macromolecules. Smaller, more focused and functionally-rich
libraries may be generated by using in methods which incorporate
predictive frameworks for hypothesis-driven protein engineering. New
design strategies and technical advances have begun a departure from
traditional protocols, such as directed evolution, which represents the
most effective strategy for identifying top-performing candidates in
focused libraries. Whole-gene library synthesis is replacing shuffling
and mutagenesis protocols for library preparation. Also highly specific
low throughput screening assays are increasingly applied in place of
monumental screening and selection efforts of millions of candidates.
Together, these developments are poised to take protein engineering
beyond directed evolution and towards practical, more efficient
strategies for tailoring biocatalysts.
Screening and selection techniques
Once a protein has undergone directed evolution, ration design or
semi-ration design, the libraries of mutant proteins must be screened to
determine which mutants show enhanced properties. Phage display methods
are one option for screening proteins. This method involves the fusion
of genes encoding the variant polypeptides with phage coat protein
genes. Protein variants expressed on phage surfaces are selected by
binding with immobilized targets in vitro. Phages with selected protein
variants are then amplified in bacteria, followed by the identification
of positive clones by enzyme linked immunosorbent assay. These selected
phages are then subjected to DNA sequencing.
Cell surface display systems can also be utilized to screen
mutant polypeptide libraries. The library mutant genes are incorporated
into expression vectors which are then transformed into appropriate host
cells. These host cells are subjected to further high throughput
screening methods to identify the cells with desired phenotypes.
Cell free display systems have been developed to exploit in vitro
protein translation or cell free translation. These methods include
mRNA display, ribosome display, covalent and non covalent DNA display,
and in vitro compartmentalization.
Enzyme engineering
Enzyme engineering is the application of modifying an enzyme's
structure (and, thus, its function) or modifying the catalytic activity
of isolated enzymes to produce new metabolites, to allow new (catalyzed) pathways for reactions to occur, or to convert from certain compounds into others (biotransformation). These products are useful as chemicals, pharmaceuticals, fuel, food, or agricultural additives.
An enzyme reactor consists of a vessel containing a reactional medium that is used to
perform a desired conversion by enzymatic means. Enzymes used in this
process are free in the solution.
Examples of engineered proteins
Computing methods have been used to design a protein with a novel fold, such as Top7, and sensors for unnatural molecules. The engineering of fusion proteins has yielded rilonacept, a pharmaceutical that has secured Food and Drug Administration (FDA) approval for treating cryopyrin-associated periodic syndrome.
Another computing method, IPRO, successfully engineered the switching of cofactor specificity of Candida boidinii xylose reductase. Iterative Protein Redesign and Optimization (IPRO) redesigns proteins to increase or give specificity to native or novel substrates and cofactors.
This is done by repeatedly randomly perturbing the structure of the
proteins around specified design positions, identifying the lowest
energy combination of rotamers,
and determining whether the new design has a lower binding energy than
prior ones. The iterative nature of this process allows IPRO to make
additive mutations to a protein sequence that collectively improve the
specificity toward desired substrates and/or cofactors.
Computation-aided design has also been used to engineer complex properties of a highly ordered nano-protein assembly. A protein cage, E. coli bacterioferritin (EcBfr), which naturally shows
structural instability and an incomplete self-assembly behavior by
populating two oligomerization states, is the model protein in this
study. Through computational analysis and comparison to its homologs, it has been found that this protein has a smaller-than-average dimeric interface
on its two-fold symmetry axis due mainly to the existence of an
interfacial water pocket centered on two water-bridged asparagine
residues. To investigate the possibility of engineering EcBfr for
modified structural stability, a semi-empirical computational method is
used to virtually explore the energy differences of the 480 possible
mutants at the dimeric interface relative to the wild type EcBfr. This computational study also converges on the water-bridged asparagines. Replacing these two asparagines with hydrophobic amino acids results in proteins that fold into alpha-helical
monomers and assemble into cages as evidenced by circular dichroism and
transmission electron microscopy. Both thermal and chemical
denaturation confirm that, all redesigned proteins, in agreement with
the calculations, possess increased stability. One of the three
mutations shifts the population in favor of the higher order
oligomerization state in solution as shown by both size exclusion
chromatography and native gel electrophoresis.
A in silico method, PoreDesigner, was developed to redesign bacterial channel protein (OmpF) to reduce
its 1 nm pore size to any desired sub-nm dimension. Transport
experiments on the narrowest designed pores revealed complete salt
rejection when assembled in biomimetic block-polymer matrices.
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