In vitro site specific mutagenesis
Objectives:
1. To understand a range of purposes for conducting in vitro mutagenesis.
2. To understand a range of phenotypes that can be sought in mutants to clarify function of a protein.
3. To understand vector types most appropriate to different kinds of in vitro mutagenesis experiments.
Background
Site specific in vitro mutagenesis is the introduction of a specific preconceived mutation. This
could be done to test a specific hypothesis about a DNA regulatory sequence, or a protein sequence, or an
RNA sequence.
Constraints on the vector.
The mutated sequence will be subjected to an assay that may require it to be in some specific
context within a vector. It is possible to conduct the mutagenesis in one vector, and then transfer the
altered sequence to another vector more specifically designed for the assay. However, in vitro
mutagenesis studies inevitably expand into a list of required mutations. It saves a lot of trouble to design
the vector so that the mutagenesis is done directly in the final vector. Furthermore, it pays to anticipate
alternative assays that may be conducted further ahead in the project and build those capabilities into the
same vector.
Promoter/Enhancer elements.
Generally, the targeted element will need to be in the context of a functional promoter and set up
do drive a reporter gene in an expression vector. If the vector will be assayed in a host other than E. coli,
it will generally need both an E. coli ori and selectable marker, and an ori for the targeted host. This is
because after mutagenesis, the clone will be grown up for sequence confirmation in E. coli, and then
transferred into the appropriate host. If the assay is to be in vitro transcription, a convenient restriction
site should be downstream to linearize the template. Other types of regulatory sites (splice sites, ori's,
recombination sites, etc.) will similarly have to be within a context wherein they can be assayed.
Promoter/enhancer regions are usually first characterized by a series of deletion mutants to localize
a region for more detailed studies. Historically, one would take advantage of available restriction sites to
make a nested set of deletions. PCR now makes it possible to form deletions starting at any arbitrary
position.
Then a finer survey of the implicated region would be done with some combination of the
following: footprinting, motif searching, gel mobility shift assays, saturation mutagenesis, and linker
scanning mutagenesis. Linker scanning mutagenesis is a systematic method of making a series of
mutants where successive regions of about 10 bases are substituted by an arbitrary sequence (usually a
restriction site).
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Linker Scanning. After McKnight and Kingsbury, Science 217:316-324 (1982).
Today, because of the reduced cost of making oligonucleotides, most people would make the 10
bp substitutions one at a time by the methods illustrated below. However, they often still call it linker
scanning.
Finally, given that a specific motif was implicated by the above methods, one might make single
base changes at the conserved position of the motif, monitoring changes in gel mobility shift, a footprint,
and expression in coordination.
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Proteins that recognize a nucleic acid sequence.
Proteins that recognize a nucleic acid sequence provide the possibility of mutating both the protein
and the binding sequence. Successful prediction of a mutation in one that compensates a mutation in the
other is generally considered as a powerful indication that one's model of the specific contacts that enforce
specificity is accurate.
Ribozymes
One of the newest targets of mutagenesis are ribozymes, RNA molecules with catalytic activity.
Ribozymes would generally be produced by in vitro transcription. If they are to be delivered in vivo, then
a transcription terminator will probably be required. Since making ribozymes is an extensive design
problem, some sort of evolutionary strategy is often used (several rounds of saturation mutagenesis
interspersed with selection for improved function).
Other RNAs
rRNA, tRNA, snRNA, etc. can be the target of in vitro mutagenesis. Because these molecules are
heavily modified, they will have to be produced in the appropriate host cells. Some provision will have to
be made to separate them from the endogenous host product. For example, one might alter their length.
Since these are usually essential genes, it will not be possible to make a null host strain, except by a
colony sectoring approach.
Signals that affect translation efficiency or half life of mRNAs may be identified by in vitro
mutagenesis.
Proteins
Mutagenized proteins for biochemical characterization will generally have to be in an expression
vector designed for mass production of the protein. The vector may also have features that aid the
purification of the protein. The promoter should be regulatable. Mutagenized proteins designed for
genetic testing will need to be in the appropriate expression vector for that target host cell. A regulatable
promoter will aid proving that the phenotypic change is really due to expression of your protein. A
regulatable promoter also helps prevent selection against your gene during construction and growth. The
promoters in many expression vectors are leaky enough that selection against the insert is a problem even
though the vector is grown under "non-expressing" conditions. Hence if the construction is complicated,
many investigators would do all the steps in some other vector, including sequence verification. Then a
restriction fragment carrying the gene would be moved to the expression vector in the last step.
One will have to provide for separation of the mutagenized protein from the endogenous protein.
Possible solutions are 1) use heterologous host, 2) delete the gene from the host, 3) use a thermostable
subject protein, or 4) use an affinity tag purification system.
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Proteins without 3-D structures
Many of the more sophisticated mutagenesis studies rely heavily on 3-D structure data. However,
there are a variety of useful experiments that can be done without 3-D information:
1. Investigating modifications (phosphorylation, glycosylation, palmitylation, etc.)
One can remove putative modification sites identified by sequence motifs to see if the resulting
protein is less modified. Further one can ask if the unmodified protein produced in this way has altered
function.
2. Investigating genetic disease:
Now that defective genes of all descriptions are being isolated by positional cloning, one is often
confronted with a defective gene with several differences from the "wild type". Mutagenesis may be
conducted to isolate each of the differences to distinguish the actual defect from inconsequential
polymorphisms. This will require an in vitro assay or a cell culture assay to act as a surrogate for the
disease phenotype. One may wish to introduce other mutations into the target gene to see how specific the
disease phenotype is with respect to different mutations. Site specific mutagenesis may also be used to
introduce the same defect into a mouse by gene replacement.
3. Identify a compartmentalization signal.
4. Assign functions to particular protein domains.
5. Delete domains for functional assignment or to stabilize for crystallography.
6. Add a tag.
7. Alanine scanning: a strategy to produce a set of mutants containing a replacement to Ala at every
position. Often this is reduced to substitution Ala at every charged residue (assuming that these have to be
on the surface). Alanine scanning is used to assign functions to different domains, and to create tentative
sets of active site residues. A variation called Cys scanning additionally allows one to covalently attach
chemical moieties at the substituted positions. Trp scanning (PNAS 92(17):7946-50 (1995)), has been
used to characterize proteins with multiple transmembrane helixes on the principle that Trp residues will
be tolerated on surfaces exposed to the lipid environment, but not elsewhere.
Alanine replacements in binding sites tend to give partial reduction in binding for reasons stated
further below. Hence, one should employ this method with a quantitative assay for the target interaction.
The loss of binding energy upon combining multiple Ala replacements at contact residues tends to be
additive. For a good introduction see Cunningham and Wells 1993. J. Mol. Biol. 234: 554-563, followed
by Lowman and Wells, 1993. J. Mol. Biol. 234:564-578.
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8. Domain addition or swapping. For example, if it is postulated that a signaling protein is activated by
regulating its access to the membrane, a constitutively active version might be made by adding a domain
that always takes it to the membrane.
9. Traditionally, active site residues have been hypothesized by sequence conservation and then altered by
in vitro mutagenesis in an attempt to define active site residues prior to obtaining 3D data. Much of this
kind of investigation has been replaced by an emphasis to get 3D structural data first, and then work out
the details of the active site by in vitro mutagenesis.
Mutagenesis of proteins with 3D structures.
Active sites
Crystal structures generally lead to a hypothesis about the roles of active site residues. However, if
a substrate analogue wasn't actually present in the crystal, the placement of substrate in the active site can
be quite speculative. Even if the enzyme was crystallized with its substrate, the resolution of a crystal
structure is generally insufficient to distinguish whether a proposed interaction (say a hydrogen bond) is
positioned well enough to contribute positively to binding energy. Therefore, the contribution of all the
proposed contacts to the activity of the enzyme still requires biochemical characterization.
One is immediately faced with a decision of what to replace these residues with. Experience has
shown that putting in a large side chain frequently kills the enzyme, leaving one to wonder if the
substrate has been completely blocked out of the active site, or if the conformation of the enzyme has been
distorted. Also, introduction of bulky side chains tend to promote aggregation either directly or by
disrupting structure and exposing hydrophobic groups. So this kind of mutant is often uninformative.
Substituting an Ala has been most consistently useful as a means of withdrawing a specific substrateenzyme contact without otherwise distorting the interaction. Gly tends to cause conformational distortion
because too much of a hole has been left in the packing of the side chains.
Other strategies are to pick chemically similar side chains to substitute, or to pick residues
appearing at this position in homologous proteins (particularly useful if the homologous protein has
altered function). One really should anticipate making a number of replacements at this position and
finding that some of them fail to fold and behave in a tractable manner. Strategies that involve changing
more than one residue at a time have an increased risk that the final product will not fold or suffer from
intractable aggregation problems..
Active site residues can be conceptualized as performing two different kinds of roles: substrate
binding, and catalysis. Mutation of catalytic residues (those that specifically stabilize the transition state)
generally kill the activity of the enzyme. One certainly has to do this experiment, because if the enzyme
still worked fine the experiment would reject the proposed mechanism. However, the negative result
(dead enzyme) is a weak result. The mutation may have killed the enzyme by altering its conformation in
some unexpected way, and the residue may in fact have nothing to do with catalysis. The overall
structural integrity of the mutant enzyme could be supported by the following observations: unaltered
proteolysis, substrate(s) still binds, other aspects of the function still carried out (eg. partial reactions not
including the mutated step), physical properties unaltered (eg. fluorescence, circular dichroism, quaternary
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structure). For small proteins (<45 kD), HSQC could both confirm minimal conformational distortion and
that the substrate still binds.
Here's an example of mutations at catalytic residues involved in the conversion of ATP + tyrosine
to tyr-AMP by tyr-tRNA synthetase. In this case there are two catalytic residues, neither one of which is
completely essential.
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Enzyme
wild type
His-45 --> Gly
Thr-40 --> Ala
His-45 --> Gly &
Thr-40 --> Ala
k(rate limiting)
38 s-1
0.16
0.0055
0.00012
KD tyr
12 uM
10
8.0
4.5
KD ATP
4.7 mM
1.2
3.8
1.1
When ATP reacts with tyrosine, the alpha phosphate must go from its normal tetrahedral
configuration through a five bonded intermediate in the transition state. This causes the other two
phosphates to be physically displaced in the active site. Thr-40 and His-45 are thought to bind the gamma
phosphate in the transition state, thus stabilizing it.
From Leatherbarrow et al. (1985) PNAS 82, 7840-7844.
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The effects of altering a substrate contact.
Mutations to binding residues normally have subtle effects on the Km, KD, or substrate specificity.
The assumption is that these residues bind the substrate the same whether or not it is in the transition state.
The substrate is usually bound by many such contacts, so disrupting one of them is insufficient to kill
activity. The binding energy contributed by the various contacts is often approximately additive.
Again looking at tyrosyl-tRNA synthetase, Thr 51 was proposed as a hydrogen bonding contact for
the ribose of the ATP.
Tyrosyl-AMP in the active site of tyrosyl-tRNA synthetase from Fersht et al., Bioch 24, 5858
The numbers tabulated below are relative Gibbs Free energies of binding determined from a
combination of equilibrium dialysis and kinetic measurements. A number of different measurements
could be compared in this way. KM values are popular to compare, because the measured KM value is
independent of enzyme concentration. This provides relief from the concern that the mutant protein is
unstable and really exists as an equilibrium between some amount that is active and some that is not. In a
simple Michaelis-Menton treatment, the KM is the same as the equilibrium dissociation constant for
binding of the substrate. However, many enzymes do not exhibit simple Michaelis-Menton behavior, and
treating them this way may cause you to attribute a mutant's effect to substrate binding when it is really
affects other aspects of enzyme structure or function. A thorough kinetic treatment (as was done here)
yields the true substrate dissociation constant, and requires actually determining how much of the enzyme
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is active, and having a detailed description of the enzyme mechanism. A final alternative is to directly
measure the substrate dissociation constant under conditions where the substrate is not turned over. This
also requires knowing how much of the enzyme is active (at substrate binding), but has the advantage that
it can be applied to mutants that are catalytically dead (at some other step besides substrate binding).
Kinetic effects of various changes to Thr-51
Enzyme
wild type (Thr-51)
Cys-51
Ala-51
Ser-51
Gly-51
kcat(s-1)
KM (mM ATP)
8.35
12.4
8.75
1.88
6.0
1.08
0.35
0.54
1.16
1.25
delta G
-0.90
-0.44
+0.92
+0.28
From Fersht and Wilkinson (1985) Bioch. 24, 5858-5861. The tabulated delta G values are computed
from kcat and KM for ATP and represent the binding energy in the transition state.
Here we see an advantage of making the comparisons as delta G values. These values are additive,
and can be broken down to individual contributions of different atoms in the side chains. If you just look
at thr -> gly, you would see that you lost .28 kcal/mol in binding energy, and you might (mistakenly)
attribute that to the loss of the hydrogen bond to the substrate. However, you really removed a hydrogen
bond, and potential interactions with 2 carbon groups (beta and gamma). The change from thr -> ser
shows that you lose -0.92 kcal/mol by withdrawing the contact with the gamma methyl group. The
change from ser -> ala shows that you actually gain 1.36 kcal/mol from withdrawing the hydrogen bond,
and the change from ala -> glycine shows that you lose another 0.72 kcal/mol from withdrawing the beta
methyl group. So thr 51 really contributes favorable hydrophobic contacts plus an unfavorable hydrogen
bond.
group
gamma methyl
gamma SH
gamma OH
beta methyl
contribution to binding energy
(kcal/mol)
-0.92
-0.46
+1.36
-0.72
A negative number denotes improved binding. Note that these numbers are in the range you
would expect for individual hydrogen bonds and Van der Waals contacts. The hydroxyl forms an
unfavorable H-bond because the distance is 0.5 angstroms too long. The unfavorable bonding energy
means that this thr residue forms a 1.36 Kcal better bond to water than the substrate, therefore it disfavors
substrate binding. Sulfhydrals tend to form longer H-bonds, therefore the cys residue makes a better
contact from this position. You could not get this information by gazing at the crystal structure, because
X-ray structures are not accurate to 0.5 angstrom resolution.
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Note that Cys actually makes a better contact with the substrate than the wild type residue, thr. If
you look at the velocity versus [ATP] curves for these two enzymes, you see that each has a range of
[ATP] where the velocity is greater than the other. This serves to show that tighter binding is not
necessarily better.
Figure modified from Fersht and Wilkinson Biochemistry 24: 585-5861 (1985)
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Mutations that alter structure.
Mutations that alter structure often have non additive effects on activity when coupled with other
mutations.
In the example of Thr 51 given above, you would expect a proline at residue 51 to contribute no
binding energy. Unexpectedly, pro 51 actually improves binding by -1.9 kcal. This is an example of a
conformational effect. Pro 51 improves substrate binding by disrupting an alpha helix and making a better
contact for His 48. This is shown by observing non additivity in double mutant Thr -> Pro 51, His -> Gly
48 (panel d below). Panels b and c show that there is not much of a conformational effect of pro 51 on
binding by Cys 35, or between mutations at residue 48 and 35.
From Carter et al. (1984) Cell 38, 835-840.
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Characterization of conformation.
In the context of active site investigation given above, a conformational effect usually arises as an
unanticipated side effect of a site directed mutation. Our current understanding of protein structure is
generally insufficient to build a preconceived conformational change into a protein. However, after the
fact, a conformational mutant can yield considerable information about structure [Biochem. 33(15):458593], stability [Biochem. 32(39):10371-7, 1993], function [J. Biol. Chem. 269(15):11578-83], or folding
[Biochem. 32(49):13566-74 (1993)]. Those who intentionally want to seek out conformational mutants
might try inserting prolines, or conducting saturation mutagenesis and screening for temperature
sensitivity.
Characterization of quaternary structure.
Refs: Jones et al., (1985) Bioch. 25, 5852-5857.
Ward et al. (1986) JBC 261, 9576-9578.
The interface between enzyme subunits can be disrupted by placing charged residues in it. The
following is an example where complementary changes are made at an interface to convert a homodimer
into a heterodimer.
Tyrosyl-tRNA synthetase is a dimeric enzyme that shows half-of-the-site reactivity. That is,
although there are two active sites, they interact with negative cooperativity such that only one is active at
a time. The experiment is to set up a situation where one can maneuver a mutation into only one of the
two subunits and thereby measure the effect on the other. In this case to physically demonstrate the
heterodimer, one of the subunits will be a version carrying a deletion to alter its size.
By engineering a negatively charged residue into the subunit interface, it was possible to cause
dissociation under native conditions by varying the pH. By engineering a positively charged residue into
the subunit interface of another variant, it was possible to form heterodimers wherein the two residues
formed a salt bridge. This verified that the pH effect was due to subunit dissociation and not due to a
generalized unfolding of the enzyme. Further, other mutations could be introduced
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specifically into one of the subunits of the heterodimer.
Mutagenesis to add a label to a protein.
Physical-chemical studies of proteins often make use of fluorescence of endogenous trp residues to
report on the conformation of the protein. With site-specific mutagenesis, one might arrange for the
placement of trp residues in ideal positions for the anticipated study. This may involve both removing
endogenous trp residues and adding new ones. (JBC 269(11):7919-25, 1994).
Other kinds of reporter groups may be added to the protein. One relatively flexible strategy is to
add a Cys at a predetermined position, and to chemically modify the Cys to attach some other moiety.
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Study Questions
1. It is proposed that the following network of hydrogen bonds is involved in stabilizing substrate binding
in a particular enzyme. How could you test this by in vitro mutagenesis?
substrate--HO Ser118 main chain NH--OH Thr46
2. What would be a good method if you wanted to map monoclonal antibody epitopes on a protein?
3. A protein has 5 trp residues. You try to change 4 to Ala so that the 5th can be used as a probe of
conformational change. This results in a protein that aggregates intractably. How might you approach
solving this problem?
4. A particular 200 bp promoter fragment is sufficient to confer a heat shock response on a reporter gene.
Can you think of a reason why a complete set of linker scanning mutants might fail to identify the heat
shock response element?
Last revised 3/13/2005 - Steve Hardies
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