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Lec. Molecular pharming: Plant production of vaccines, antibodies,
therapeutic proteins- transient expression of recombinant proteins in plant
using viral vectors.
Molecular farming is the production of pharmaceutically important and commercially
valuable proteins in plants. Its purpose is to provide a safe and inexpensive means for the mass
production of recombinant pharmaceutical proteins. Complex mammalian proteins can be
produced in transformed plants or transformed plant suspension cells. Plants are suitable for the
production of pharmaceutical proteins on a field scale because the expressed proteins are
functional and almost indistinguishable from their mammalian counterparts.
Biopharmaceuticals While early biopharmaceuticals were mainly proteins for replacement
therapy such as erythropoietin (EPO) or growth hormones, current blockbusters include
monoclonal antibodies (mAbs) or other proteins for the treatment of cancer and inflammatory
diseases. Sales of many top-selling biopharmaceuticals are increasing,with US sales of
themonoclonal antibody Herceptin used for the treatment of breast cancer growing by 82% from
2005 to 2006 alone (Lawrence, 2007). In 2007, the three top-selling pharmaceutical products sold
by the Swiss drug-maker Roche were therapeutic monoclonal antibodies (Rituxan/Mabthera,
Herceptin, and Avastin), contributing to sales of 9311 million € (Roche, Annual Finance Report for
2007).
The microbial system like E.coli is commonly used for production of recombinant protein
production. The main drawback of the E.coli system is that it cannot be used for the production of
glycoprotein. Simple proteins that are not glycosylated (such as insulin and human growth
hormone) or that, while glycosylated in their native form, do not require glycosylation for
pharmacological activity (such as α-, β-, and γ-interferon (IF), interleukin-2 (IL-2)) are mainly
produced in E. coli.
Apart from E.coli, some of the yeast like Pichia pastoris and Saccharomyces is also used
for the production of recombinant protein. The main advantage is the it can produce glycoprotein
and the yeast system is free from toxin that is associated with E.coli system. But some of the
glycoprotein produced is immunogenic.
Most of the human protein are glycoprotein and needs glycosylation for their proper
function. Mammalian cell culture system is commonly used for the production of
glycoprotein. For the production of more complex proteins requiring posttranslational
modifications (like glycosylation –addition of carbohydrate moiety to the protein) the
Chinese hamster ovary (CHO) cell is the main workhorse used. Further mammalian cell
types used for the commercial production of approved protein drugs are baby hamster
kidney (BHK) cells, mouse myeloma cell lines. The main advantage of this is the
glycoprotein produced is not immunogenic in human or animal. However, this is a very
expensive system for production of recombinant protein. Both the microbial and
mammalian cell culture system is expensive process that require huge capital investment,
trained main power to operate the system and operation cost is high thereby increasing
the cost of the recombinant protein produced. Plant is a cheap alternative for the
production of the recombinant protein. It requires less capital investment and scale up can
be easily done in case of requirement. The main drawback of the plant system is that the
glycoprotein produced are immunogenic and purification of the recombinant protein form
the plant tissue following good laboratory practices is difficult. However in large number
of recombinant proteins have been successfully expressed in the plant and some are in
clinical trial before commercialization. It is estimated that the plant derived recombinant
protein can be 10-20 fold less expensive than the fermenter based recombinant protein
production. Also the initial investment for establishment is less and scale up is possible.
Molecular Pharming — plants as production hosts for high-value compounds
Plant cells combine the advantage of a full posttranslational modification potential with simple
growth requirements and basically unlimited scalability ofwhole plants in the field. Plants and plant
cells are versatile production systems, also allowing targeting of the recombinant proteins
produced to different organs or subcellular compartments, thus allowing improved protection
against proteolysis. Plants are not known to harborhuman or zoonotic pathogens,making thema
safe host for the production of biopharmaceuticals. Human growth hormone was the first
recombinant protein with therapeutic potential that was successfully expressed in plants (Barta et
al., 1986). More complex proteins soon followed, and Hiatt et al. (1989) produced a correctly
assembled complete IgG1 antibody with full binding functionality in tobacco plants. Over the
following years, a vast variety of pharmaceutical proteins were expressed in different plant species
(with tobacco remaining the main production host) and targeted to different organelles or organs.
Some examples include the expression of human serum albumin in tobacco and potato (Sijmons
et al., 1990), human α-IF in rice (Zhu et al., 1994), and, more recently, IL-12 in tomato (GutiérrezOrtega et al., 2005). A milestone was the successful production of recombinant secretory IgA
(sIgA) in tobacco plants (Maet al., 1995). It is believed that such sIgAs have great potential for
use in topical passive immunotherapy (Fischer and Emans, 2000).
Examples of whole-plant-derived biopharmaceuticals in preclinical and clinical development
The first clinical trial of a plant-produced biopharmaceutical was conducted by Planet
Biotechnology Inc. for CaroRx™ (also known as the secretory variant of Guy's 13), a secretory IgA
against Streptococcus mutans, which is the main causative agent of tooth decay. CaroRx is
produced in tobacco grown in the field. The antibody was shown to effectively eliminate
recolonization of S. mutans in humans for at least 4 months after oral topical application (Ma et al.,
1998). CaroRx has received approval for human use in the E.U. (Kaiser, 2008). Large amounts of
CaroRx are required for topical applications and it is dosed at 22.5mgper course of treatment (Ma
et al., 1998). If itwere administered to the child population in Europe alone, more than 1000 kg of
the antibody would be required annually (Ma et al., 2005). Another company which is expecting to
enter a plant-produced protein product into the market soon is Ventria Bioscience, which uses its
ExpressTec technology for high-level production of recombinant proteins in field-grown rice. Two
of Ventria's primary products are human lactoferrin and human lysozyme. Expression levels
reported are extremely high, with the recombinant protein contributing to 60% of total soluble
protein or 1% of rice flour weight (10 g/kg) in the case of lysozyme and to 25% of total soluble
protein or 0.5% of rice flour weight (5 g/kg) in the case of lactoferrin (Huang, 2005). The company
is developing several products containing human lactoferrin and human lysozyme. An oral
rehydration solution containing both enzymes as a dietary supplement for acute pediatric diarrhea
is expected to be released soon.
Plants are genetically modified to produce pharmaceuticals, also called plant-made
pharmaceuticals (PMPs). The first recombinant plant-derived pharmaceutical protein (PDP) was
human serum albumin, initially produced in 1990 in transgenic tobacco and potato plants. Fifteen
years on, the first technical proteins produced in transgenic plants are on the market, and proof of
concept has been established for the production of many therapeutic proteins, including antibodies,
blood proteins, cytokines, growth factors, hormones, recombinant enzymes and human and
veterinary vaccines.
Plant production of vaccines
Vaccinology is a rapidly expanding research field and new vaccination
strategies have been
developed thanks to modern technologies based on the rational design of attenuated pathogens,
live recombinant vaccines and
protein (antigen)- or peptide (epitope)-based subunit vaccines
(Plotkin
2005). The aim of these approaches is to obtain efficient and safe vaccine
formulations being able at the same time to induce effective, long lasting
immunity against
complex viral pathogens such as HIV-1 (Nabel 2001), influenza A/H5N1 virus (Stephenson et al.
2004), SARS-coronavirus
(Stadler and Rappuoli 2005). To reach this goal, targets must be
found able not only to generate long-term memory B cell producing neutralizing antibodies to
block free pathogens, but also T cell mediated immunity for the control of pathogen spreading by
the elimination of infected cells
(Lambert et al. 2005). The
possibility to produce subunit
vaccines through plants paves new ways and
offers solutions to some of the problems
associated to traditional production systems.
Protein antigens from various pathogens have been expressed in plants and used to
produce immune responses resulting in protection against diseases in humans. Plantderived vaccines have been produced against Vibrio cholerae, enterotoxigenic E. coli,
hepatitis B virus, Norwalk virus, rabies virus, human cytomegalovirus, rotavirus, and
respiratory syncytial virus F. Edible vaccines may be particularly valuable as a low-cost delivery
mechanism for immunization against various diseases in developing countries. In 1992, Mason et
al. (1992) described the expression of Hepatitis B virus (HBV) surface antigen in tobacco, paving
the way not only for the production of antigens for parenteral application in plants, but also for
the production of “edible vaccines”. The HBV surface antigen was also expressed in transgenic
lupin and lettuce plants. When mice and human volunteers ate the antigen containing plants
they developed a specific antibody response (Kapusta et al., 1999). Other plants considered for
oral vaccine delivery are tomato, carrot, banana, and corn. Medicinal and commercial utility of
vaccines for human use produced in plants will depend on the performance in human clinical
trials. According to Marshall (2007), several vaccines, including oral vaccines, were in clinical
trials (or had completed clinical trials) in October 2007, conducted by the group of C. Arntzen at
Arizona State University and by companies such as Large Scale Biology Corporation and
Prodigene. Human safety testing has successfully been completed for orally delivered spinach
expressing a rabies antigen (Yusibov et al., 2002), E. coli heat-labile toxin as a vaccine for
diarrhea produced in transgenic maize (ProdiGene Inc.) and transgenic potato tubers (Tacket
et al., 1998), Hepatitis B virus surface antigen in transgenic potato (Thanavala et al., 2005) and
transgenic lettuce (Kapusta et al., 1999) and Norwalk virus capsid protein in transgenic potato
(Tacket et al., 2000). There is a concern that edible vaccines might induce tolerance instead
of immunity, thereby actually weakening an immune response to the antigen, although signs of
such induced tolerance have not been reported yet (Vermij, 2004).
Several antigens have been successfully expressed in plants by stable nuclear transformation
with different ends. In some cases, plants were
considered only as “biofactories” for massive
production, while in others edible plant varieties have been chosen for the direct oral delivery of
the expressed antigens.
Plant production of antibodies
Antibodies are multi-subunit glycoproteins, produced by the vertebrate immune system. They
recognize and bind to their target antigens with great affinity and specificity, which allows them to
be used for many applications, including the diagnosis, prevention and treatment of human and
animal disease. It is estimated that approximately 1000 therapeutic recombinant antibodies are
under development, up to one quarter of which may already be undergoing clinical trials. A large
proportion of these antibodies recognize cancer antigens but others have been developed for the
diagnosis and treatment of infectious diseases, acquired disorders and even transplant rejection
(Gavilondo and Larrick 2000). Most of the recombinant full-length immunoglobulins being
developed as
pharmaceuticals are produced in mammalian cell culture, a few in hybridoma
lines but most in immortalized lines which have been cleared by the
FDA and equivalent
authorities in other countries.
Over the last 15 years, plants have emerged as a convenient, economical and scalable alternative
to the mainstream antibody production systems which are based
on the large-scale culture of
microbes or animal cells (Chu and Robinson 2001; Wurm 2004). In this chapter, we discuss the
advantages and disadvantages
of plants for antibody production, the diverse plant-based
systems which are now available, and factors governing the success of antibody production in
plants.
Recombinant antibody technology Structure of natural antibodies
The typical antibody format is the mammalian serum antibody, which
comprises two identical
heavy chains and two identical light chains joined by disulfide bonds (Figure 1).
Fig. 1. Structure of a typical mammalian serum antibody, comprising two identical heavy chains
(dark gray) and two identical light chains (light gray). Solid black lines indicate continuation of the
polypeptide backbone (simple lines indicate the constant parts of the antibody, curly lines indicate
the variable regions). Antibody domains are indicated by circles. Disulfide bonds are represented
by thick black bars.
Each heavy chain is folded into four domains, two either side of a flexible hinge region which
allows the multimeric protein to adopt its characteristic Y-shape. Each light chain is folded into
two domains. The N-terminal domain of each of the four chains is variable, i.e. it differs among
individual B cells due to unique rearrangements of the germ-line immunoglobulin genes. This
part of the molecule is responsible for antigen recognition and
binding. The remainder of the
antibody comprises a series of constant domains, which are involved in effector functions such
as immune cell recognition and complement fixation. Below the hinge, in what is known as the
Fc portion of the antibody, the constant domains are class-specific.
Mammals produce five
classes of immunoglobulins (IgG, IgM, IgA, IgD and IgE) with different effector functions. The Fc
region also contains a conserved asparagine residue at position 297 to which N-glycan chains
are added. The glycan chains play an important role both in the folding of the protein and the
performance of effector functions. Antibodies are also found in mucosal secretions, and these
secretory antibodies have a more complex structure than serum antibodies. They are dimers
of the serum-type antibody, the two monomers being attached by
an additional component
called the joining chain. There is also a further
polypeptide called the secretory component,
which protects the antibodies from proteases (Figure 2).
Fig. 2. Structure of a mammalian secretory antibody, comprising a dimer of the typical serum
antibody, and including two additional components, the joining chain (black disc) and secretory
component (white disc). Heavy chains are shown in dark gray and light chains in light gray. Solid
black lines indicate continuation of the polypeptide backbone (simple lines indicate the constant
parts of the antibody, curly lines indicate the variable regions). Antibody domains are indicated
by circles. Disulfide bonds are represented by thick black bars.
Production of secretory IgA in transgenic plants.
The assembly and expression of the multimeric, complex molecule secretory
immunoglobulin (sIgA) was first described successfully in transgenic plants and plants
remain the best system to express this molecule. Indeed, the large scale production of
recombinant sIgA is a very challenging task for tow main reasons: (i) the components of
this molecule are naturally produced by two distinct cell types (plasma and epithelial
cells) and (ii) the final product is a large complex molecule of almost 400 kDa display
numerous post-transcriptional modifications (intra and inter chain disulfide bonds and
glycosylation sites). SigA has also been produced in CHO cells but the cost of production
might bee too high to envisage commercialization.
Antibodies
The secretory IgA consists of two IgA molecules linked by a joining (J) chain and
associated with a secretory component (SC). To exert its protective activity on the
mucosa, polymeric IgAs (pIgAs, mostly IgA dimmers) are transported across the
epithelium after binding to the polymeric immunoglobulin receptor (pIgR), which is
expressed basolaterally on the epithelial cells. During transport, the pIgR (also known as
the transmembrane secretory component) is cleaved and the secretory component (SC) is
released in association with pIgA to form sIgA. Quantitatively, sIgA is the most
important antibody class with 40- 60 mg/ kg produced every day, whereas the daily
production of IgG is only of 30 mg/kg. Moreover, the covalent binding of SC enhances
resistance to proteolytic degradation making their the most stable form of Ab. In humans,
there are two IgA subclasses, IgA1 and IgA2, that differ only in the hinge region: IgA1
contains a 13 amino acid, proline rich sequence which is not present in IgA2. The amino
acid composition of the IgA hinge region renders it more resistant to proteases then other
immunoglobulins. However, IgA1 is particularly sensitive to proteases produced by
Gram-negative bacteria, whereas IgA2 is relatively more resistant (sue to the absence of
the proline rich region)
The J chain is 137 amino acid, 15.6 kDa glycoprotein that is added just before the
secretion of pIgA by the plasma cells. The J chain is a key protein in the synthesis of
sIgA because it promotes polymerization of IgA and because its presence in the polymer
is required for their affinity of pIgR/secretory component.
The secretory component (SC), an 80 kDa glycoprotein is the extracellular domain of the
pIgR synthesized by the mucosal epithelium. During basal-to-apical transport across the
epithelial cells, the pIgR ectoplasmic domain is cleaved, releasing SC in association with
pIgA, thus forming sIGA. The human pIgR contains 5 domain. Domain I of the pIgR is
involved in the initial non-covalent attached to pIgA. Subsequently, during transcytosis,
the pIgR becomes covalently attached to pIgA via cysteine residues in domain V.
Passive immunization with sIgA
Active immunization has been successful in protecting against several infectious
diseases. However, vaccines are still not available for numerous pathogens (eg. Human
immuno deficiency virus, respiratory syncytial virus, hepatitis virus). Furthermore,
active immunization is generally less effective in immuno-compromised individuals.
Passive immunization with sIgA should provide a better protection level against
pathogens than monomeric antibodies (i.e. IgGs or IgAs). It has been shown that SC is
essential for the stability of the whole sIgA molecule when targeting the gastrointestinal
(GI) tract.
Furthermore, sIgAs have a higher binding avidity; of r their antigens than monomeric
antibodies, because of their four antigen combining sites.
Production of recombinant sIgA.
Thus, for only two-expression system have been successful in producing rec. sIgA.
1. Full length sIgA was first expressed in transgenic tobacco
2. In the past 5-6 years, several groups have succeeded in expressing recombinant
sIgA in mammalian cells. The use of CHO (Chinese hamster ovary) cells to
produce sIgA has been the most successful strategy, and has relied on the
successive stable transfecion and selection of CHO clones capable of expressing
monomeric, dimeric and finally sIgA recombinant molecules. However, the levels
of antibody production is stable transfectomas have generally been lower than in
murine hybridomas.
Production of recombinant sIgA in plants.
Production of full length antibodies in plants.
Plants have the ability to assemble immunoglobulin heavy chains and light chains
to form full-length antibodies very efficiently. In plants, the assembly of immunoglobulin
takes place in ER. Signal peptide sequences directs translocation into the lumen of the
endoplasmic reticulum (ER). Both plant and non-plant signal sequences form a variety of
sources are sufficient for ER-targeting. Plant chaperones homologous to mammalian BiP,
GRP94 and PDI (Protein disulfide isomerase) have been described within the ER, and the
expression of immunoglobulin chains in plants is indeed associated with increased BiP
and PDI expression. Co-expression of heavy and light chains resulted in IgG assembly
and displacement of BiP from the heavy chain as the amount of light chain increased.
Murine IgG1 (Guy’s 13) that binds to the adhesion protein of Streptococcus mutants, the
primary cause of dental caries. The strategy used to produce this antibody in plants was to
express each immunoglobulin chain separately in different plant lines and then to stack
the two genes in the same plant line by crossing parental plants individually expressing
the heavy and light chains. This involved two generation of plants, and using this
technique, the yield of recombinant antibody was consistently high (approximately 1%
TSP). Guy’s 13 IgG is relatively easy to purify in large quantities from tobacco. Guy’s 13
can also be expressed in transgenic plants with a transmembrane sequence so that it is
retained in the plasma membrane with variable regions protruding into the apoplasm.
Transgenic plants expressing antibodies immobilized in such a fashion may have
important applications in phytoremediation and phytomining.
Other groups have expressed IgG antibodies using double transformation technique or
have cloned the light and heavy chain genes together in a single Agrobacterium T-DNA
vector. Both strategies can save time and effort.
Production of multimeric antibodies: sIgA.
The ability to stack gene in transgenic plants by successive crosses between
individually transformed parental plants is a considerable advantage in attempting to
construct multimeric protein complexes, such as secretory antibodies.
In order to generate a secretory antibody version of Guy’s 13 in plants the carboxyl
terminal domains of the Guy’s 13 IgG antibody heavy chain were modified by replacing
the Cγ3 domain with the Cα2 and Cα3 domain of an IgA antibody, these being required
for binding to the J chain and SC. Four transgenic plants were generated to express
independently the Guy’s 13 kappa chain, the hybrid IgA-G antibody heavy chain, the
mouse J chain and the rabbit SC. A series of sexual crosses was performed between these
plants and filial recombinants in order to generate plants in which all four protein chains
were expressed simultaneously. In the final, quadruple transgenic plant, three froms of
the antibody was detectable by western blot anlysis of samples prepared under nonreducing conditions. These bands had approximate molecular mass of 210 kDa
(monomeric IgA-G), 400 kDa (IgA-G dimerized with the J chain) and 470 kDa (dimeric
IgA-G associated with the SC). The assembly was very efficient, with greater than 50%
of the SC being associated with dimeric IgA-G.
Frigerio et al. demonstrated that secretion of sIgA-G proceeded at a very slow rate in
tobacco leaf cells. After 24 hour only about 10% of newly assembled molecules had been
secreted with the bulk probably remaining in the ER.
Finally, in a human trial, the plant-derived secretory Guy’s 13 antibody prevented oral
colonization by Streptococcus mutants, thereby demonstrating for the first time the
therapeutic application in humans of a recombinant product derived from plant.
The Guy’s 13 sIgA-G plantibody technology is licensed to Planet Biotechnology Inc.
(USA) and is currently in clinical trials under the product name CaroRxTM.
Transient transformation To get round the difficulties encountered in obtaining high rate of
expression of the heterologous gene in nuclear transformed plants, transient transformation
strategies have been devised. In this case, the gene encoding the antigen of interest is inserted
in the genome of a pathogen that is used as a vector for expression during plant infection. The
two major techniques used to get transient transformation are based on plant viruses- or
Agrobacterium-mediated infection of fully developed plants.
Plant viral expression vectors for recombinant protein expression
The transgenic approach has some shortcomings, including the length of time required to
obtain the transgenic producer lines, low levels of expression, and inherent difficulties in
the modification of an existing product. In some expression hosts, scaling up production
also takes a ling time.
The foreign gene is inserted into the viral genome so that, upon infection of the host plant
cell, the transgene is replicated and expressed along with native viral genes. This method
of transient expression adds several further advantages to plant-based expression,
including improved time efficiency, higher levels of target protein expression, flexibility
and convenience in the modification of existing products (or the development of new
ones), ease of scale-up, flexibility in the selection of a production host and the potential
for protein manufacture in contained facilities. Furthermore, target proteins (in particular
antigenic peptides) can be genetically fused to viral structural proteins, such as coat
proteins, so that the plant virus is used not only for expression but also for the delivery of
the vaccine antigens. To date, the coat proteins from a number of plant RNA viruses have
been successfully used as carrier for antigenic peptides derived from various pathogens.
Plant viral vectors are being successfully developed and exploited for the industrial-scale
expression of heterologous proteins and as a research tool for studies of gene expression. The
initial engineering strategy (the ‘full virus’ vector strategy) aimed to design a vector that was
essentially a wildtype virus, which was modified to carry and express a heterologous sequence
that encoded a gene of interest. The new emerging trend (the ‘deconstructed virus’ vector
strategy) reflects an ideology that recognises the inherent limitations of the viral process. It
attempts to ‘deconstruct’ the virus, by eliminating functions that are limiting or undesired, and to
rebuild it, either by delegating the missing necessary functions to the host (which is genetically
modified to provide those functions) or by replacing them with analogous functions that are not
derived from a virus.
Plant RNA viruses as expression vectors
The majority of viruses that infect plants have single-stranded, positive sense
RNA genomes. It has therefore been necessary to use infectious cDNA clones for the in
vitro manipulation of RNA viruses, allowing them to be developed as effective tools for
the commercial production of target protein in plants. Siegel conceptualized the potential
use of RNA viruses as expression vectors. Brome Mosaic Virus (BMV) and Tobacco
Mosaic Virus (TMV) were the first two RNA viruses to be converted into expression
vectors. The genomes of number of plant RNA viruses have been engineered to express
target sequences. Based on genome structure and identification of virus gene functions,
several approaches have been employed for the expression of foreign sequences using
plant viruses as expression systems.
These include:
(i)
replacing non-essential viral genes with target sequences.
(ii)
Inserting target sequences into the viral genome as an additional gene with
an additional promoter.
(iii) Fusing target sequences with viral structural gene with a cleavage site or
read-through sequence.
(iv)
Functional complementation of defective viral components, and
(v)
Trans-complementation of viral genes through transgene expression in the
host plant.
Examples of plant viruses used in the development of expression vectors
Virus
Strategies used
Pathogen or protein
epitope
Tobacco Mosaic
CP replacement, second subgenomic Malaria, rabies virus
Virus
promoter (sgp), CP fusion, Read
FMDV, MAB, alphathrough fusion with CP, fusion with trichosanthin (Inhibitor of
cleavage site
HIV), allergens, scFvs,
HCV(Hepatitis C virus
vaccine)
Potato Virus X
Second sgp, CP fusion, CP fusion
GFP, rotavirus, scFv
with FMDV 2A element, IRES
elements.
Cowpea Mosaic
Fusion with CP, fusion to CP with
GFP, HIV
Virus
protease cleavage site,
complementation
Alfalfa Mosaic Virus Fusion with CP, second sgp
HIV, rabies, Measles,
RSV
RNA Virus + and - strand
In RNA viruses, the viral RNA may be directly replicated or act as a template in the
synthesis of DNA. The RNA of single-stranded RNA viruses may be the positive strand
(the mRNA or the negative strand (complementary of the mRNA). RNA viruses with (-)
strand and dsRNA genomes must carry the RNA polymerase in the viral particle because
the incoming viral RNA into plant cell can be neither translated nor copied by the cellular
machinery. Deprotinized RNA genome (only nucleic acid) are not non-infectious. In
contrast (+) strand viral particles lack a virion polymerase and deprotinized RNA (only
viral RNA) are infectious. All RNA viruses except retroviruses encode an RNAdependent RNA polymerase (Replicase) to catalyze the synthesis of new genomes and
mRNA. (The intact virus particle, which is referred to as a virion consists of a nucleic
acid molecule encased by a protein capsid. In some of the more complex virions, the
capsid is surrounded by a lipid bilayer and glycoprotein containing envelope, which is
derived from a host cell membrane).
Tobacco Mosaic Virus
TMV is (+) RNA virus. It causes leaf mottling and discoloration. TMV is a rodshaped particle. Its ~2130 identical copies of coat protein subunits (158 amino acid
residue; 17.5 kDa) are arranged. TMVs single RNA strand (~6400 nt).
Tobacco Mosaic virus (TMV) genomic and subgenomic RNAs.
A. 6400 nucleotide TMV genomic RNA acts as a messenger RNA for the expression
of the two replicase-associated proteins of 126kDa and 183 kDa.
B. The other genes in TMV RNA are expressed form subgenomic RNAs (sgRNA)
during the replication cycle.
The rod shaped TMV virions are composed of the CP protein and the genomic RNA.
5’end of TMV RNA contain a typical 5’ terminal cap structure (m7 GpppG) was
discovered in TMV RNA. Infectivity lost on removal of cap structure. The 3’ terminal
five codons of the 180 kDa protein gene overlap with the third ORF, coding for the 30
kDa protein. This third ORF terminates two nucleotides before the initiation codon of the
fourth ORFs which encodes CP. The genes located internally in the genome are not
expressed from the genomic RNA, but from subgenomic RNAs, which are generated
during the course of replication.
The viral genome codes for two replicase-associated proteins translated right from the
genomic RNA, and a movement protein (MP), and a CP translated from 3’ coterminal
subgenomic mRNAs. The genome of TMV employs two distinct strategies for protein
expression: read-through of an amber stop codon and production of subgenomic RNAs.
The 5’-ORF of TMV is translated from the genomic RNA resulting in the accumulation
of a 126 kDa protein. A translational read-through of an amber stop codon results in
synthesis of a 183 kDa protein instead of the 126 kDa protein. These two proteins are
first translated from the genomic RNA and comprise the viral RNAdependent RNA
polymerase (RdRp) synthesizing a fulllength minus-sense copy of the genome. These
copies are then used as a template for the amplification of plus sense genomes and the
subgenomic mRNAs for MP and CP synthesis. The MP, a 30 kDa protein, is produced
early in the infection cycle and is required for cell to- cell movement of progeny viral
RNA genomes by modifying the size exclusion limit of plasmodesmatal junctions. Later
in infection the production of genomic RNA, CP mRNA, and CP become the major
events in the viral replication cycle. The CP, a 17.5 kDa structural protein produced at
very high levels, is required for systemic movement of the virus through the vascular
system in the plant and for encapsidation of the viral genome.
Two general expression vector designs for TMV based vectors were developed, CP
fusion vectors and dual subgenomic expression vectors.
1. CP Fusion vector
In CP fusion vectors, a ‘‘read-through sequence’’ is introduced immediately after the CP
stop codon and before the ORF of interest. Peptides of interest, e.g., the epitope of the
murine hepatitis virus, are fused to the CP and are therefore incorporated into soluble
virions presenting the peptide on the surface of the virion. TMV particles have also been
used as an epitope-presentation system. The helical arrangement of 2130 copies of coat
protein around the viral RNA allows the presentation of multiple copies of the foreign
epitope on the virus surface. At the same time, this can be detrimental to virion stability
because the extra peptide can destabilize the virion structure. To circumvent this problem,
a more stable system was developed exploiting the read through sequence from the
replicase gene, so that both wild type and fusion peptide containing coat proteins were
produced.
2. Subgenomic expression vectors
A schematic diagram of the TMV-based dual subgenomic vector described herein
is shown in Fig. 1. This 30B-based hybrid vector is designed on the characteristics
identified by a series of TMV expression vectors
Invitro production of infective recombinant TMV-RNA and inoculation of N.
benthamiana plants.
Capped infectios viral RNA is generated in vitro from the linearised plasmid with the
ribomax Large Scale RNA production system-T7 kit (Promega) and the Cap-Analog
(Ambion) according to manufacture’s instructions.
T7 RibomaxTM Express Large scale RNA production system is an in vitro transcription
system designed for the consistent production of milligram amounts of RNA in a short
amount of time. DNA templates are usually linearized prior to in vitro transcription to
produce RNA transcripts of defined length. Blunt end obtained by treating with DNA
polymerase I Large (Klenow fragment)
T7 reaction components:
Ribomax Express T7 2X Buffer
Linear DNA template (1 µg total)
Nuclease Free water
Enzyme Mix, T7 Express
Sample reaction
10 µl
1-8 µl
0-7 µl
2 µl
--------Final volume
20 µl
--------Mix gently and incubate at 37 C for 30 min.
rNTPs. Enzyme Mix: T7 RNA polymerase, recombinant RNasin- Ribonuclease inhibitor
and Rec. inorganic pyrophosphatase.
Cap Analog (Ambion)
- m7 G(5’) ppp (5’) G (Cap Analog) is used for the synthesis of 5’ capped RNA
molecules in vitro transcription reactions. Capped mRNAs are genrally translated
more efficiently in reticulocyte lysate and wheat germ in vitro translation systems.
Uncapped mRNAs are rapidly degraded after microinjection into cells.
Infection of tobacco plant with TMV RNA
The integrity of the transcribed RNA is assessed by electrophoresis on a 1% agarose gel.
After dusting N. benthamiana leaflets (2 per plant) lightly with carborundum powder, 2
μg in vitro transcripts are gently rubbed on their surface to initiate infection.
Recombinant protein isolation
Between one and three weeks after inoculation, leaflets of infected plants are frozen in
liquid nitrogen and pulverized to isolate the recombinant protein in PBS buffer.
TMV expression vector was used for the expression of subunit vaccine against hepatitis
C virus (HCV). A consenses sequence matching hypervariable region 1 (HVR1) of HCV,
encoding a potential neutralizing epitope of 27 amino acids was fused to the C-terminus
of CTB. Mice immunized intranasally with plant extract containing ~0.5 – 1.0 µg
CTB/HVR1 developed anti-HVR1 antibodies. The same epitope has been engineered as
fusion with other plant viruses such as AIMV. Human volunteers (in FDA approved
trials) fed with spinach containing recombinant particles generated both IgG and IgA
response specific to the pathogen.
Due to size limitations, there is unstable long distance movement of the chimeric virus in
tobacco.
Transfection of N. benthamiana plants
About four-week-old N. benthamiana plants approximately at a 10-leaf stage are suitable
hosts for transfection with agrobacteria to subsequently produce viral RNA. Veins of two
leaves per plant are wounded carefully by pricking with a needle. Stem and leaves of the
plant are then submerged for twenty minutes into the overnight culture of Agrobacterium.
The N. benthamiana plants treated in that way are then placed in the dark in a humid
atmosphere for 24 h to recover from the treatment. Thereafter, plants are kept in a growth
chamber under 16 h of daylight at 22 C.
TMV vectors have been used to produce of many different kinds of proteins in plants including
allergens [5,6], antibodies [7] or antibody fragments [8], and vaccine candidates [9,10]. TMV is an
RNA virus that expresses large amounts of coat protein from a viral subgenomic promoter. To
convert TMV to an expression vector an additional, heterologous coat protein subgenomic
promoter and restriction enzyme sites for cloning of foreign DNA sequences were inserted into a
T7 promoter driven cDNA clone of TMV[11]. In vitro transcription of this plasmid with T7 RNA
polymerase is needed to generate biologically active transcripts. Transcripts are typically rubinoculated by hand onto plants to initiate an infection [4]. The in vitro transcription and rub
inoculation steps in particular, add significantly to the cost and complexity of using TMV vectors.
Agroinfection [12,13] is a less-expensive and more reproducible strategy for infecting plants with
RNA viruses. In agroinfection a plant-functional promoter and RNA virus cDNA are transferred as
T-DNA from Agrobacterium tumefaciens into plant cells. The T-DNA is transcribed in planta, to
generate biologically active viral RNAs that can initiate self-replication. Although agroinfection has
been used for several different plant RNA viruses, it has not been routinely used with TMV-based
vectors. Recently, an agroinfection-compatible TMV replicon was constructed with extensive
modifications to the TMV cDNA. These modifications included (1) the deletion of the TMV coat
protein (CP) gene, (2) generating nearly 100 point mutations to destroy cryptic introns in the viral
cDNA, and (3) inserting multiple (up to 24) plant-gene introns into the TMV cDNA sequences in a
binary vector [14]. The point mutations and inserted introns dramatically improved the infectivity
of the TMV replicon delivered to plants by agroinfection. However, because of the deletion of the
CP gene, this replicon cannot move systemically in plants. Plants were therefore infected in a
process called "magnifection" [9,15] in which whole plants were submerged and infiltrated with A.
tumefaciens cultures carrying intron-modified TMV sequences in a binary vector.
The ‘full virus’ strategy Under the ‘full virus’ scenario, the delivery of amplicons is achieved by
infecting the host with a mature viral particle or with a DNA/RNA molecule that contains a
complete copy of the viral vector. Depending on the vector and its ability to move systemically, 2–
3 weeks will be necessary for the vector to move to as many organs and cells of the host plant as
possible. For example, the ability of viruses to infect the host is naturally low and typically requires
either mechanical injury of the host tissues or delivery by an insect vector. Furthermore, systemic
spread is usually a very species-specific process that is easily impaired by genetic manipulation
of the virus. The full-virus strategy by designing vectors that were capable of systemic spread.
The limitations of this approach are obvious: inserts larger than 1 kb are often not expressed
properly and do not move well systemically; only short epitopes (of 20 amino acids or less) can be
presented effectively as fusions to coat protein; systemic vectors never infect all of the
harvestable parts of the plant (e.g. they rarely infect lower leaves); the process is asynchronous
as it progresses at different speeds in different leaves; and the vector is usually unstable, thus
much of the infected tissue does not express the protein of interest. These of vectors based on
the ‘full virus’ strategy seems to be constrained, primarily by the limitations on the number and
type of proteins that can be expressed using this technology. The most obvious application for
these vectors today seems to be for the production of vaccines as coat protein fusions of short
antigenic peptides or through the chemical coupling of antigens to engineered viral particles.
The ‘deconstructed virus’ approach may rely on the use of Agrobacterium to efficiently
introduce a DNA copy of the viral vector into a plant cell (Table 1). All of the processes necessary
for the generation of a functional DNA or RNA amplicon from T-DNA occur in plant cells. In fact,
‘agroinfection’ has been used for many years because it is often much more efficient than infection
using assembled viruses, and definitely much more efficient than the use of DNA or RNA
infectious molecules. In case of RNA viruses, agroinfection also represents an inexpensive
alternative to the in vitro transcription methods that are used to convert a DNA vector into an
infectious RNA. The agrodelivery of the two genomic components of cowpea mosaic virus from a
mixed suspension of bacteria cultures, each harbouring different subgenomic complements, was
demonstrated in an interesting recent publication. Two elements of first-generation viral vectors
that are limiting are the ability of the vector to move systemically, which is provided by the coat
protein, and the low level of expression of the protein of interest, probably because a significant
fraction of the cell metabolic resources are devoted to synthesis of a large amount of coat protein.
A simple solution was to eliminate the coat protein and replace the systemic movement ability by
artificial delivery of the viral vector to the entire plant using Agrobacterium.
On the basis of these findings, a simple fully scalable protocol for heterologous protein
expression in plants was designed that is devoid of stable genetic transformation of a plant, but
instead relies on the transient amplification of viral vectors delivered to the entire plant using
Agrobacterium. The process is in essence an infiltration of whole mature plants or of detached
mature leaves with a highly diluted suspension of agrobacteria carrying a proviral replicon on the
T-DNA. In this case, infiltration of agrobacteria replaces the conventional viral functions of primary
infection and systemic movement (Figure 1). Amplification within each cell and movement from
cell-to-cell is performed by the replicon. Depending on the vector used, the host organism and the
initial density of bacteria, the process takes from 4 to 10 days and, depending on the specific
gene of interest, can result in the expression of up to 5 g recombinant protein per kg of fresh leaf
biomass or over 50% of total soluble protein. Furthermore, as the viral vector does not contain a
coat protein gene, it can express longer genes (up to 2.3 kb inserts or up to 80 kDa proteins). The
infiltration of plants/detached leaves with bacteria has been achieved in many ways, one simple
process being vacuum infiltration after immersing the aerial part of the plant in a bacterial
suspension and applying a weak vacuum (approximately –0.8–0.9 bar) for 10–30 s [13__,14,15].
This new expression strategy, called magnifection (Figure 2), combines the advantages of three
biological systems: the speed and expression level/yield of the virus, the transfection efficiency of
Agrobacterium, and the posttranslational capabilities and low production cost of plants.
Schematic description of infection and spread of replicons based on (a) first-generation and
(b) second-generation viral vectors.
General scheme for recombinant protein production in plants using magnifection.
Depending on the protein of interest, magnifection can produce up to 0.5–5 g recombinant
protein per kg of leaf biomass. With a green biomass yield of 100–300 ton/hectare per year, the
yield of recombinant protein is expected to be 50–600 kg/hectare/year. The expression time is 6–
10 days, making magnifection especially attractive for those applications where rapid industrial
manufacturing is required. Components (a), (e) and (f) are standard existing industrial processes.
Components (b), (c) and (d) are new elements, because large-scale infiltration (b) requires special
equipment and the use of bacteria in steps (b), (c) and (d) requires biological containment to
prevent the release of genetically engineered bacteria into the open environment.
Industrialization of the magnifection technology is currently an essential part of the activity of
several business and academic groups, and the most serious constraint is the need for a
specialized facility that allows handling of plants treated with agrobacteria and that can operate
under conditions compliant with current good manufacturing practice. TMV is an RNA virus that
expresses large amounts of coat protein from a viral subgenomic promoter. To convert TMV to an
expression vector an additional, heterologous coat protein subgenomic promoter and restriction
enzyme sites for cloning of foreign DNA sequences were inserted into a T7 promoter driven cDNA
clone of TMV[11]. In vitro transcription of this plasmid with T7 RNA polymerase is needed to
generate biologically active transcripts. Transcripts are typically rub-inoculated by hand onto
plants to initiate an infection [4]. The in vitro transcription and rub inoculation steps in particular,
add significantly to the cost and complexity of using TMV vectors. Agroinfection [12,13] is a lessexpensive and more reproducible strategy for infecting plants with RNA viruses. In agroinfection a
plant-functional promoter and RNA virus cDNA are transferred as T-DNA from Agrobacterium
tumefaciens into plant cells. The T-DNA is transcribed in planta, to generate biologically active
viral RNAs that can initiate self-replication. Although agroinfection has been used for several
different plant RNA viruses, it has not been routinely used with TMV-based vectors. Recently, an
agroinfection-compatible TMV replicon was constructed with extensive modifications to the TMV
cDNA. These modifications included (1) the deletion of the TMV coat protein (CP) gene, (2)
generating nearly 100 point mutations to destroy cryptic introns in the viral cDNA, and (3)
inserting multiple (up to 24) plant-gene introns into the TMV cDNA sequences in a binary vector
[14]. The point mutations and inserted introns dramatically improved the infectivity of the TMV
replicon delivered to plants by agroinfection. However, because of the deletion of the CP gene,
this replicon cannot move systemically in plants. Plants were therefore infected in a process called
"magnifection" [9,15] in which whole plants were submerged and infiltrated with A. tumefaciens
cultures carrying intron-modified TMV sequences in a binary vector. While the magnifection
process is very efficient, it is not easily adapted to a high throughput workflow. Also, the increased
size of the intron-modified vectors can make cloning into these vectors more challenging. In
addition, it is not clear if the intron-modified vectors are absolutely required for efficient
agroinfection of local and systemic infection of plant tissue with a TMV vector. Maps
TMV is a rod-shaped virus that has a single-stranded plus-sense RNA genome. TMV expresses
four proteins from three open reading frames (ORFs). Two viral genes (the viral movement protein
and the capsid protein) are expressed from separate subgenomic promoters. TMV has typically
been modified to express foreign genes by either replacing a viral gene (such as the coat protein
[CP] gene, for example) with a gene of interest (for review, see Scholthof et al., 1996) or by
inserting an additional subgenomic promoter (Dawson et al., 1989; Donson et al., 1991; Pogue et
al., 1998) into the viral genome to drive the expression of an inserted foreign gene. Plants can be
inoculated with TMV vectors through a process called agroinfection. In agroinfection, A.
tumefaciens was used to deliver T-DNA composed of 35S promoter-driven TMV cDNA to plant
cells. Transcription of T-DNA in the plant nucleus generated RNA that was capable of initiating
self-replication in the cytoplasm. Multiple reports have documented the low agroinfection
efficiency of the typical 35S-driven TMV vector (Turpen et al., 1993; Marillonnet et al., 2005; Man
and Epel, 2006). Here, we report on the construction of an improved agroinfection-compatible
TMV vector that lacks the TMV CP gene coding sequence. This modification resulted in a vector
with several significant improvements, such as (1)much higher agroinfection efficiency; (2) higher
recombinant protein expression levels; and (3) inability to form virus particles during its infection/
replication cycle. This new expression vector is called the TMV RNA-based overexpression
(TRBO) vector. Here, we demonstrate that the TRBO vector can produce up to 100 times more
recombinant protein than the P19-enhanced agroinfiltration transient expression system
described above. It is proposed that, because of its efficacy and ease of use, the TRBO vector will
be a useful transient expression vector for production of recombinant proteins in plants for either
research or production purposes.
Figure 1. Maps of plasmids used in this project. The T-DNA regions of binary plasmids used in this
project are represented. Block arrow, CaMV duplicated 35S promoter. Black box, CaMV polyA
signal sequence/ terminator. Dark gray box, Tobacco etch virus 5#-nontranslated leader
sequence. Light gray box, Ribozyme. Bent arrows, Subgenomic promoters. ORFs are represented
by white boxes. Identities of ORFs are labeled in white boxes. Replicase, TMV 126K/183K ORF;
MP, movement protein; P19, 19-kD RNA-silencing suppressor gene from Tomato bushy stunt
virus.
Another advantage of TRBO is that it does not produce TMV CP. Because TMV CP is required
for systemic movement, TRBO is not capable of systemic movement in plants. It also will not
produce virions in plants. This has definitive biocontainment and protein purification advantages.
First, this feature reduces the chances for inadvertent plant-to-plant movement of the vector.
Second, when extracting proteins from TRBO-infected tissue, the recombinant protein of interest
does not need to be purified away from virion particles. If one is using a viral vector that does
generate virus particles (such as JL24), efforts must be taken to both separate virion particles
from the recombinant protein of interest and also to inactivate virus particles in any extracts of
infected plant materials. These issues are not a concern with TRBO because it does not generate
virus particles.