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Chapter 11
Protein Engineering
Engineering Disulfide Bonds
improving Stability in Other Ways
Changing Binding Site Specificity
Structural Scaffolds
Directed Evolution
Adding New Functional Groups Using Nonnatural Amino
Recombinant Domains
DNA Shuffling
Combinatorial Protein Libraries
Biomaterials Design Relies on Protein Engineering
Engineered Binding Proteins
Overlapping-extension PCR
Increasing the range of industrial enzymes has three facets.
First, modem biology has identified many novel enzymecatalyzed reactions that may be of industrial use.
Second, it is now possible to produce desired proteins in large
amounts because of gene cloning and expression systems.
Third, the sequence of the protein itself may be altered by genetic
engineering to improve its properties. This is known as protein
Disulfide bonds are especially important for those proteins found outside the
cell in oxidizing environments.
In practice, most enzymes used industrially will be exposed to
such oxidizing conditions, and therefore disulfide bonds are particularly relevant.
Introduction of extra disulfide bonds is a relatively straightforward way to increase the
stability of proteins.
The first step is to simply introduce two cysteine residues into the polypeptide chain.
Then, under oxidizing conditions, these will form a disulfide bond provided that the
polypeptide chain folds so as to bring the two cysteines into dose contact.
Obviously, for this approach to work, the tertiary structure of the protein must be
known so that the cysteines can be inserted in appropriate positions (Fig. 11.1).
In general, the longer the loop of amino acids between the two cysteines, the greater the
increase in stability.
Note however, that formation of a disulfide linkage can create a strained conformation if
the two cysteines are not properly aligned. This may result in
a decrease in stability of the protein.
Introduction of Disulfide Bonds
A disulfide bond can be added to a protein by changing two amino acids into cysteines by site-directed
mutagenesis. When the engineered protein is put under oxidizing conditions, the two cysteines form a 10
disulfide bond, holding the protein together at that site.
This approach has been demonstrated using the
bacterial virus T4.
The polypeptide chain of 164 amino acids folds into two domains and has
two cysteines, neither involved in disulfide bond formation in the wild-type
One of these, Cys54, was first mutated to Thr to avoid formation of
incorrect disulfides.
The other, Cys97, was retained for use in disulfide formation. Extensive
analysis of possible locations for disulfides was carried out. Those disulfides
that might impair other stabilizing interactions in the protein were
This left three possible disulfide bonds that should theoretically promote
stability, located between positions 3 and 97, 9. and 164, and 21 and 142
(Fig. 11.2).
Disulfide Engineering of T4 Lysozyme
T4 lysozyme has two domains. The N-terminal region is shown in green and red, and the C-terminal
region is in blue. These are linked by an alpha helix (purple). Disulfide bonds were added at three
locations in T4 lysozyme to increase the stability. The first disulfide was between positions 9 and 164.
This links the first alpha helix at the N terminus with the C-terminal tail. The second disulfide is
between positions 2 and 97, which links the N- and C-terminal domains. Finally, the third disulfide links
position 21 in the N-terminal domain with 142 in the C-terminal domain. The figure depicts alpha
helices as barrel-shaped and beta-sheets as green arrows.
To test these experimentally, five amino adds
(Ile3, Ile9, Thr21,
Thr142, and Leu164) were converted to Cys in various
Stability was measured by thermal denaturation; the melting temperature,
Tm, is the temperature at which 50% of the protein is denatured.
Because of the effects of entropy, the greater the number of possible unfolded
conformations, the more likely a protein is to unfold. Decreasing the number of possible
unfolded conformations therefore promotes stability.
disulfide linkages.
Glycine (-)
proline (+)
hydrophobic residues
Because of its asymmetrical structure, the alpha helix is actually a dipole with
a slight positive charge at its N-terminal end and a slight negative charge at its Cterminal end. The presence of amino add residues with the corresponding opposite
charge dose to the ends of an alpha helix promotes stability. In natural proteins the
majority of alpha helixes are stabilized in this manner. However, in cases where such
stabilizing residues are absent, protein engineering may create them.
Asparagine and glutamine residues are relatively unstable. High
temperature or extremes of pH convert these amides to their corresponding adds,
aspartic add and glutamic add. The replacement of the neutral amide by the
negatively charged carboxyl may damage the structure or activity of the protein. This
may be avoided by engineering proteins to replace Asn or Gln by an uncharged
hydrophilic residue of comparable size, such as Thr.
In addition to altering the overall stability of a protein, it is possible to
deliberately change the active site.
The most straightforward alterations to make are those that change the binding
specificity for the substrate
enzyme mechanism.
or a cofactor, but do not disrupt the
Changing the specificity for a cofactor or substrate may be useful, either to make the
product of the enzyme reaction less costly or to change it chemically.
This principle has been demonstrated with several enzymes that use the cofactors
NAD or NADP to carry out dehydrogenation reactions.
NAD is used by dehydrogenases in
degradative pathways, and the respiratory chain oxidizes the resulting
NADH. In contrast, biosynthetic enzymes use NADP. Structurally
they differ only in NADP having an extra phosphate group attached to the ribose ring
(Fig. 11.3). This gives NADP an extra negative charge and, not surprisingly, enzymes
that prefer NADP have somewhat larger binding pockets with positively charged
amino acid residues at the bottom. Enzymes that favor NADH often have a
negatively charged amino acid residue in the corresponding position.
Difference in Structure between NAD and NADP
NAD (nicotinamide adenine dinucleotide) differs from NADP by one single phosphate (yellow)
Several enzymes that use NAD or NADP have been engineered to change their reference
For lactate dehydrogenase (LDH) of most bacteria uses reduced NAD, not
NADP, to convert pyruvate to lactate.
A conserved aspartate provides the negative charge at the bottom of the cofactor
binding pocket that excludes
If this is changed to a neutral residue, such as serine, the enzyme becomes able to
use both NAD and NADP. If, in addition, a nearby hydrophobic residue in the cofactor
pocket is replaced by a positively charged amino
arginine), the enzyme now prefers NADP to NAD (Fig. 11.4).
acid (such as lysine or
Changing Cofactor Preference of Lactate Dehydrogenase
Lactate dehydrogenase (LDH) preferentially binds NAD because the binding pocket has an aspartic
acid. The negatively charged carboxyl repels the negatively charged phosphate of NADP. Changing
the aspartic acid to serine allows either NAD or NADP bind to LDH. Adding a positively charged lysine
makes the pocket more attractive to the NADP.
LDH for its substrate
The specificity of
can be altered in a
similar way. The natural substrate lactate is a three carbon hydroxyacid. It is
possible to alter several residues surrounding the substrate binding site
without impairing the enzyme reaction mechanism.
By replacing a pair of alanines with glycines, the binding site can be made
larger. By replacing hydrophlic residues (Lys, Gln) with hydrophobic ones (Val, Met), the
site becomes more hydrophobic. Alteration of multiple residues gives an engineered
LDH that accommodates five or six carbon analogs of lactate and uses them as
Relatively few of the amino acid residues in a
protein are actually involved in the active site.
Most of the protein provides the 3D platform or
scaffold needed to correctly position the active
site residues. Quite often the scaffold is much
larger than really necessary.
For example, the β-galactosidase of
Escherichia coli (LacZ protein) has
approximately 1000 amino acids, whereas
most simple hydrolytic enzymes have only 200
to 300. Presumably it should be
possible to redesign a functional βgalactosidase that is only 25% to 30% the
size of LacZ protein. From an industrial
viewpoint, such a smaller protein would
obviously be more efficient.
Directed evolution is a powerful technique to alter the function of an
enzyme without the need for exhaustive structural and functional data. Directed
evolution can be used to change substrate specificity, either changing the enzyme to
recognize a totally different substrate, or making subtle changes where the substrate is
slightly different. The main premise of directed evolution is the random mutagenesis
of the gene of interest, followed by a selection scheme for the new desired function.
Directed evolution screens for new enzyme activities by constructing a library of different
enzymes derived from the same original protein. Each protein in the library
is slightly different because of random mutagenesis. Random mutants may be
generated over the entire length of the gene sequence. Alternatively, certain target
amino acids can be replaced
by random amino acids. The third
main method for generating mutants relies on recombination
(homologous or nonhomologous).
These mutant genes are then screened for the new, desired enzyme activity after
insertion into a suitable expression vector and host cell.
Random mutagenesis usually starts with a gene whose function is dose to that desired.
The gene is randomly mutated throughout the entire sequence using error-prone
PCR (epPCR). Different methods exist to induce errors during PCR amplification. The
most straightforward is to add MnC1, to the PCR reaction. Taq polymerase has a fairly
high rate of incorporating the wrong nucleotide, and MnCI, stabilizes the mismatched
bases. The error will be copied in subsequent rounds of amplification. Adding nucleotide
analogs such as 8-oxo-dGTP and dITP, which form mismatches on the opposite strand,
can also enhance the PCR error rate. These analogs in combination with MnCl2, can
induce a wide variety of different mutations along the length of a gene. Some random
mutations that occur outside the active site may cause subtle changes with profound
effects on substrate recognition and enzyme function.
The third method to form. new enzymes via directed evolution involves various schemes
for recombining different domains. These are based on homologous or nonhomologous
sequences and encompass a variety of different protocols, including
DNA shuffling and
combinatorial protein libraries.
High conc.
Many nonnatural amino adds have different functional groups that are useful in protein
engineering. For example, adding p-benzoyl-L-phenylalanine( pBpa) into one position
of glutathione-S-transferase (GST) adds a crosslinking group that can be activated by
UV irradiation. When GST is modified in this way, UV irradiation creates a covalently
linked homodimer (Fig. 11.6).
Adding New Functional Groups to Proteins
(A) Nonnatural amino acids add new functional groups. These can be incorporated into a protein
during translation. (B) The nonnatural amino acid, pBpa, crosslinks the GST mutant protein to form a
Incorporating a nonnatural amino acid into a protein can be done on a small scale
by isolating the tRNA for a particular amino acid and attaching the nonnatural amino
acid. The charged tRNA is then added to an
in vitro protein translation
system, which incorporates the nonnatural amino acid into the growing polypeptide
chain. The chemical method is too costly and time consuming for large-scale use.
For large-scale incorporation, E. coli may be modified to insert the nonnatural amino
add in vivo. Inserting a nonnatural amino acid during in vivo protein synthesis requires
mutant aminoacyl-tRNA synthetase
that charges a
tRNA with the nonnatural amino add. The laboratory of Peter G. Schultz, at the Scripps
Research Institute, has developed an E. coli strain that incorporates pBpa at a specific
amber codon (One of the three terminator codons.
Its sequence is UAG ).
Directed evolution was used to mutate a tyrosyl-tRNA synthetase from M. jannaschii.
The enzyme from M. jannaschii was used because it does not recognize any
endogenous E. coli tRNA. Consequently, it needs the gene for its specific partner tRNA
to be provided as well. In addition, the partner tRNA was altered so that it recognizes
the amber stop codon instead of its original natural codon (i.e., it is an amber mutant
tRNA). The result is that when the mutant tyrosyl-tRNA synthetase is expressed in27 E.
coli, it inserts whichever amino acid it is charged with at amber stop codons.
To alter the tyrosyl-tRNA synthetase, a library of mutant enzymes was generated
by random mutation of each amino acid residue involved in recognition of the amino
acid substrate (originally tyrosine). The library of mutant tRNA synthetase genes
was transformed into E. coli that possess a gene for the partner tRNA, and a gene for
chloramphenicol resistance with an amber codon in the middle. The
E. coli were grown in the presence of pBpa and chloramphenicol. If the mutant tRNA
synthetase was able to insert an amino acid at the amber stop codon, then the
chloramphenicol resistance gene (CAT) was expressed, and the cells lived.
Otherwise, the cells died. This was the positive selection (Fig. 11.7).
This positive selection does not exclude mutant tRNA synthetases that
charge the amber tRNA with a natural amino acid, so a negative
selection scheme was used next. The plasmids carrying the mutant tRNA
synthetases were isolated and transformed into a different E, coli strain. This
toxin gene with an amber suppressor
mutation plus the amber tRNA. Here the mutants were grown without
E. coli had a
any pBpa. If the mutant tRNA synthetase could charge the amber tRNA with
a natural amino add, the toxin would be made and the E. coli would die. This
eliminated mutant tRNA synthetases that used natural amino adds. The
selection scheme was repeated numerous times, and finally a specific mutant
tRNA synthetase was isolated (Fig. 11.7) that recognized the amber tRNA
and linked pBpa to it.
Positive and Negative Selections for Mutant tRNA Synthetase
toxin gene
chloramphenicol resistance gene (CAT)
Positive and Negative Selections for Mutant tRNA Synthetase
Tyrosyl-tRNA synthetase normally attaches tyrosine to the tRNA for the CUA amber codon. The amino acids
that recognize tyrosine were randomly mutagenized to form a library of different tRNA synthetases that still
recognize the same tRNA, but might attach different amino acids. Next, these library clones (MutTyrRS) were
expressed in a cell containing another plasmid carrying genes for the amber tRNA (Mj tRNACUA) and for
chloramphenicol acetyl transferase (CAT). In the middle of the CAT gene is an amber codon (amberCAT).
These E. coli are grown with pBpa and chloramphenicol. MutTyrRS must charge the Mj tRNACUA with pBpa or
another amino acid to express CAT, which protects the E. coli from chloramphenicol. The library clones that
survive this selection are expressed in a different E. coli host (left). This strain has the gene for the amber
tRNA (Mj tRNACUA) plus a toxin gene with an amber codon. The toxin protein is made if the MutTyrRS charges
the amber tRNA with any amino acid (no pBpa is present). This eliminates clones that charge the amber tRNA
with an endogenous amino acid. The positive and negative selection schemes are alternated to find the best
mutant tRNA synthetase.
A related approach to directed evolution is to deliberately recombine
functional domains from different proteins.
An example is the creation of novel restriction enzymes by linking the
cleavage domain from the restriction enzyme FokI with different sequencespecific DNA binding domains.
FokI is a type II restriction
with distinct N-terminal and C-terminal domains that function
in DNA recognition and DNA cutting, respectively.
By itself the endonudease domain cuts DNA nonspecifically. However, when
the nuclease domain is attached to a DNA-binding domain, this domain
determines the sequence specificity of the hybrid protein. The two domains
may be joined via a sequence encoding a linker peptide such as
(GlyGlyGlyGlySer), (Fig. 11.8). Cleavage of the DNA occurs several bases to
the side of the recognition sequence, as in the native FoKI restriction enzyme.
Recombining Domains to Create a Novel Endonuclease
The FokI endonuclease has separate nuclease and sequence recognition domains. Using genetic
engineering, the recognition domain of FokI can be replaced with a Gal4 recognition domain, which
binds to a different DNA sequence. The two domains are joined with an artificial linker peptide. The
new hybrid enzyme now cuts DNA at different locations from the original FokI protein.
Several different DNA binding domains have been combined with the
nuclease domain of FokI.
For example, the Gal4 protein of yeast is a transcriptional activator that
recognizes a 17 base-pain consensus sequence. Gal4 has two domains, a
DNA binding domain and a transcription activating domain. The N-terminal
147 amino acids of Gal4 can be fused to the nuclease domain of FokI, giving
a hybrid protein that binds to the Gal4 consensus sequence and cleaves the
DNA at that location.
Zinc finger domains have also been joined to the nuclease
domain of FokI. The zinc finger is a common DNA binding motif found in
many regulatory proteins. The zinc finger consists of 25 to 30 amino adds
arranged around a Zn ion, which is held in place by binding to conserved
cysteines and histidines. Each zinc finger motif binds three base pairs,
and a zinc finger domain may possess several motifs. In this example,
domains approximately 90 amino adds long and comprising three zinc
finger motifs, and which therefore specifically recognize nine base DNA
sequences, were connected to the FokI nuclease domain (Fig. 11.9).
Assembly of Zinc Finger Domains
(A) The nuclease domain of FokI can be linked to a zinc finger domain containing three zinc finger
motifs. Zinc fingers recognize three nucleotides each; therefore, any 9-base-pair recognition sequence
can in principle be linked to the nuclease domain. (B) The sequence of the hybrid between the FokI
nuclease and the zinc finger domain. The letters represent the amino acid sequence. The amino acids
in large letters recognize and bind the DNA sequence.
Zinc finger motif: 25 aa
genome editing
Natural selection works on new sequences generated both by
and recombination.
DNA shuffling is a method of artificial evolution that includes the
creation of novel mutations as well as recombination. The gene to be
improved is cut into random segments around 100 to 300 base pairs long.
The segments are then reassembled by using a suitable DNA polymerase
with overlapping segments or by using some version of overlap PCR. This
recombines segments from different copies of the same gene (Fig. 11.10).
Mutations may be introduced in several ways, including the standard
mutagenesis procedures. In addition, the DNA segments may be generated
using error-prone PCR instead of by using restriction enzymes. Alternatively,
mutations may be introduced during the reassembly procedure itself by
using a DNA polymerase that has impaired proofreading capability. The result
is a large number of copies of the gene, each with several mutations
scattered at random throughout its sequence. The final shuffled and mutated
gene copies must then be expressed and screened for altered properties of
the encoded protein.
FIGURE 11.10
DNA Shuffling for a Single Gene
Introducing point mutations and shuffling gene segments can generate a better version of a protein.
First, many copies of the original gene are generated with random mutations. The genes are then cut
into random segments. Last, the fragments are reassembled using overlap PCR. The new constructs37
must be assessed for enhanced protein function.
FIGURE 11.11
DNA Shuffling for Multiple Related Genes
Shuffling segments from related genes can also enhance the function of a particular protein. The
original set of related genes are digested into small fragments and reassembled using PCR. The new
combinations are tested for a change in function.
A more powerful variant of DNA shuffling is to start with several closely
related (i.e, homologous) versions of the same gene from different
organisms. The genes are cut at random with appropriate restriction
enzymes and the segments mixed before reassembly. The result is a
mixture of genes that have recombined different segments from different
original genes (Fig. 11.11). Note that the reassembled segments keep their
original natural order.
For example, several related β-lactamases from different enteric bacteria
have been shuffled. The shuffled genes were cloned onto a plasmid vector
and transformed into host bacteria. The bacteria were then screened for
resistance to selected β-lactam antibiotics. This approach yielded improved
β-lactamases that degraded certain penidins and cephalosporins more rapidly
and so made their host cells up to 500-fold more resistant to these p-lactam
Another approach to protein engineering is to generate large numbers of
different protein sequences and then screen them for some useful enzyme
activity or other chemical property. (Screening is often done by phage
display Antibody: light-chain library and heavy-chain library).
Rather than merely generating large numbers of random polypeptides,
combinatorial screening usually uses pre-made modules of some sort to
create a random shuffling library. For example,
protein motifs known to provide a binding site for metal ions or metabolites
might be combined with segments known to form structures such as an
alpha helix.
In a common approach, DNA modules of around 75 base pairs (i.e., 25
codons) are made by chemical DNA synthesis. Several modules are then
assembled to give a new artificial gene. The modules are usually joined by
PCR using overlapping primers (Chapter 4). Modules may be joined in a
chosen order or in a randomized manner. For proper expression of the
assembled sequence, the front and rear modules are normally specified to
provide suitable promoter and terminator sequences. The intervening
modules may then be randomly Generation of shuffled to generate more
possible variation (Fig. 11.12).
FIGURE 11.12
Generation of Random Shuffling Library
To create a library of new proteins, different modules can be randomly joined together. The first module
(yellow) has sequences for the promoter; therefore, this is always added at the front. Similarly, the last
module (purple) has the terminator sequences. Using overlapping PCR primers, the modules are joined
together in a particular order (part A), or randomly (part B). Because random assembly creates many
different combinations, this method creates a library of new proteins that can be screened for a
particular function or set of functions.
Combinatorial library: exon shuffling
FIGURE 11.13
Generation of Alternative Splicing Library
The exons from an original gene can be recombined such that one exon is missing in each novel
construct. The new genes are then screened for new or altered function.
Making drugs specific for a particular organ can eliminate many unwanted
side effects. One, way of achieving this is to attach the drug to a reagent
that recognizes proteins specific to particular tissues.
antibodies are the most widely used reagents for binding specific target
proteins. However, antibodies require disulfide crosslinks to function, and
these are often hard to maintain during large-scale manufacture. Some
researchers have therefore been seeking alteratives to antibodies.
To generate novel binding domains, a binding protein with a known structure
is chosen and the amino acid residues associated with binding are identified.
The binding protein is modified by mutation of these residues and then
screened for new binding partners. It is hoped that the targeted directed
evolution approach will find new, more easily isolated proteins for targeting
drugs to specific target cells within our bodies.
FIGURE 11.14
Structural Domains Involved in Protein-Protein Binding
Some types of protein backbones used as scaffolds for protein-binding agents. The proteins used (with their
Protein Data Bank ID numbers) are beta sandwich (1FNA, fibronectin); beta barrel (A chain of 1BBP, lipocalin);
thee-helix bundle (1Q2N, SpA domain); repeat proteins (1MJ0, AR protein); peptide binders (chain A of 1KWA,
PDZ domain); small scaffolds (chain F of 1MEY, zinc-finger protein); scaffolds presenting constrained peptides
(chain A of 2TrX, thioredoxin A); proteins with intrinsic fluorescence (chain A of 1GFL, GFP) or intrinsic
enzyme activity (1M40, beta-lactamase); protease inhibitors (1ECY, ecotin); and disulfide-bonded scaffolds
(chain A of 1CMR, scorpion toxin). Cysteine residues and disulfide bonds are depicted in yellow. From: Binz
and Plückthun (2005). Engineered proteins as specific binding reagents. Curr Opin Biotechnol16, 459–469.
Reprinted with permission.
Monoclonal Antibodies as Therapeutic Agents
About 100 years ago, horses were inoculated with the bacterium
Corynebacterium diphtherial
However, the administration of horse antiserum containing antibodies
against the exotoxin provides passive immunity, protecting the patient
from a fatal outcome when the antiserum is given within the first few days of
the onset of infection.
Patients often develop their own antibodies against the foreign proteins of
either whole or partially purified therapeutic antiserum.
After a second treatment, the sensitized patient may go into anaphylactic
shock and die.
Today, with the advent of hybridoma methodology (mouse antibodies),
antibodies are once again seen as potential therapeutic agents. This
technique can be used to maintain a continuous supply of pure
monospecific antibody.
Therefore, the present goal is to design and produce human
monoclonal antibodies with both specific immunotherapeutic
properties and lowered potential for immunogenicity.
Structure and Function of Antibodies
An antibody molecule
(immunoglobulin) consists of two
identical "light" (L) protein
chains and two identical "heavy"
(H) protein chains.
The N-terminal regions of the L and H
chains together form the antigen
recognition site of each antibody.
(Fig. 10.5).
The sites that recognize and bind
antigens consist of three
Figure 10.5 Structure of an
regions (CDRs) that lie within the
antibody molecule. The H and L
variable (VH and VL) regions at the
chains contain variable (VL and VH)
N-terminal ends of the two H and two
and constant (CL, CH1, CH2, and CH3)
L chains. The CDRs are the part of
domains with their CDRs (CDR1,
an antibody molecule with the
CDR2, and CDR3). The Fv, Fab, and
greatest variability in amino acid
Fc portions of an antibody molecule
are delineated. The N-terminal (NH2) sequence.
and C-terminal (COOH) ends of each
polypeptide chain are indicated.
Fc portion elicits several immunological responses after
antigen-antibody binding occurs:
The complement cascade is activated. The
components of this system break down cell membranes,
activate phagocytes, and generate signals to mobilize
other components of the immunological response system.
Antibody-dependent cell-mediated cytotoxicity
(ADCC), which is the result of the binding of the Fc
portion of the antibody to an Fc receptor of an ADCC
effector cell (natural killer cells, neutrophils,
macrophages), is produced. The bound effector cell
releases substances that lyse the foreign cell to which the
Fab portion of the antibody molecule is bound.
After the Fab region binds to a soluble antigen, the Fc
portion of an antibody can be bound to Fc receptors
of phagocytic cells, which engulf and destroy the
antibody-antigen complex.
Treating Brain Tumors
Anti-EGF receptor Ab
Human Monoclonal Antibodies
Although the initial studies of immunotherapeutic agents were promising,
there are drawbacks to chemical coupling and the use of a nonhuman
monoclonal antibody.
Finally, if the therapy requires multiple treatments, the antibody
component should be from a human source to prevent immunological
cross-reactivity and sensitization of the patient.
It is very difficult to create specific non-cross-reactive antibodies
because of problems associated with obtaining human monoclonal
antibodies by conventional hybridoma techniques.
1. The human chromosomes of fused human lymphocyte-mouse
myeloma cells are unstable, and so cells producing a human monoclonal
antibody are extremely rare.
2. No effective human myeloma cell line that can replace the mouse
myeloma cell line in the procedure has been discovered.
3. For ethical reasons, it is not acceptable to immunize humans against
various antigens.
In one scheme, human B lymphocytes that were actively producing
antibody were isolated from subjects and treated with a fluorescencelabeled antigen. Fluorescence-activated cell sorting (FACS) was then
used to enrich the cell sample for specific antibody-producing B lymphocytes.
Because B cells do not grow well in culture, they were transformed with
Epstein-Barr virus, which allows them to grow more readily in culture.
Some of these transformed B-cell clones were found to secrete human
monoclonal antibody that interacted with the selecting antigen. Unfortunately,
with this strategy, the yields tend to be small and the monoclonal
antibodies have weak antigen binding affinities. In addition, there is a
low probability that a nonimmunized individual will have antibodysecreting cells that recognize the selecting antigen.
It may also be possible to introduce cells of the human immune system into
a mutant mouse strain that lacks, for the most part, its own natural
immunological cell repertoire. After the transplantation of human immune
system stem cells, such a mouse, with severe combined
immunodeficiency (scid mouse), acquires a human cell immune system
and, in response to challenge by antigen, can produce human antibodies.
Other workers are attempting to introduce human immunoglobulin genes
into the germ line of mice to create transgenic mice that could produce a
human immunoglobulin after immunization with a particular antigen.
Standard hybridoma methodology could be used to isolate cells that
secreted a specific monoclonal antibody from the transgenic animals; then
these positive cell lines could be screened to determine which lines produced
antibody that was encoded by the human immunoglobulin transgenes. In fact,
a transgenic mouse that expresses the native form of both H and L human
immunoglobulin genes has recently been reported.
The humanization of the immune system of scid mice and the generation of
transgenic mouse lines are both laborious ways to produce human
monoclonal antibodies. Consequently, researchers have been examining
genetic engineering strategies to create both human therapeutic antibodies
and effective dual-function proteins that have the ability to bind a target and
then destroy it.
Hybrid Human-Mouse Monoclonal Antibodies
The modular nature of antibody functions has made it possible to convert a
mouse monoclonal antibody into one that has some human segments but
still retains its original antigen binding specificity. This hybrid antibody is
called a chimeric antibody, or a "humanized" antibody.
The first portion of a mouse monoclonal antibody that was targeted for
replacement with a human sequence was the mouse Fc fragment. It was
chosen because the mouse Fc fragment functions poorly as an effector of
immunological responses in humans; it is also the most likely fragment
to elicit the production of human antibodies.
When a chimeric antibody that
contained the binding site from a
mouse monoclonal antibody directed
against the surface of human colon
cancer cells was tested in patients
with colorectal cancer, it remained
in the blood system about six
times longer than the complete
mouse monoclonal antibody did,
thereby extending the period of
effectiveness. Only 1 patient of the
10 developed a mild antibody
response to the chimeric antibody.
To diminish immunogenicity and to introduce human Fc effector capabilities,
the DNA coding sequences for the Fv regions of both the L and the H
chains of a human immunoglobulin were substituted for the Fv DNA
sequences for the L and H chains from a specific mouse monoclonal
antibody (Fig. 10.9). This replacement of Fv coding regions can be
accomplished either by using oligonucleotides and in vitro DNA replication or
by using subcloned segments. The DNA constructs for both chimeric chains
were cloned into an expression vector and transfected into cultured B
lymphocytes (????) from which the chimeric antibody was collected.
The "humanizing" of mouse and rat
monoclonal antibodies has been
taken one step further than the
formation of chimeric molecules
described above by substituting into
human antibodies only the CDRs of
the rodent monoclonal antibodies
(Fig. 10.10). Because these
"reshaped" human antibodies have
antigen binding affinities similar to
those of the original rodent
monoclonal antibodies, they may be
effective therapeutic agents.
Figure 10.10 Genetically engineered humanized antibody. The CDRS
(CDR1, CDR2, and CDR3) from the genes for H and L immunoglobulin chains
of a mouse monoclonal antibody replace the CDRS of the genes for a human
antibody. The product of this constructed gene is an immunoglobulin with the
antigen binding specificity of the mouse monoclonal antibody (pink) and all
the other properties of a human antibody molecule (light blue).
Humanizing rodent monoclonal antibodies may be performed as follows.
Starting with a rodent hybridoma cell line, cDNAs for the L and H
chains can be isolated.
degenerate primers (VL and VH)
signal sequences
constant regions
The variable regions of these cDNAs can be amplified by PCR. The
oligonucleotide primers that are used for this amplification are
complementary to the sequences at the 5' and 3' ends of the DNA
encoding the variable regions, where there is a high degree of nucleotide
sequence conservation from one antibody gene to another. From the
nucleotide sequences of the cDNAs for the light and heavy regions (VL and
VH), it is possible to delineate the limits of the CDRs.
It is usually straightforward to determine where the CDRs begin and
end, since these regions are highly variable in sequence while the
sequences of the framework regions are relatively conserved.