Why would a mammalian vector plasmid require an antibacterial resistance gene?

Why would a mammalian vector plasmid require an antibacterial resistance gene?

We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

I have been trying to understand the composition of plasmids used in recombinant DNA cloning, such as this: and I am wondering what purpose the ampicillin resistance gene has when transfected into a mammalian cell host (as it says it is for).
I understand that ampicillin resistance is used for selection in bacterial cells so that only transformed bacteria are grown as well as ampicillin being needed to prevent bacterial contamination among a mammalian cell culture, but I don't see why mammalian cells require a resistance gene to an antibiotic.


I think the assumption you are making is that the ampicillin resistance gene would save a purpose in the mammalian cells. It does not.

As you said, it is only use during the cloning process to select the bacteria to replicate and produce more of it.

There is a resistance gene on the vector you posted which affect mammalian cells: neomycin. In this case, once transfected into mammalian cells, you could select only the cells with the plasmid by adding neomycin the cell culture medium.

Note: neomycin selection can take some time (4-5 days) and higher dose than for example puromycin. Hope this helps!

Molecular Cloning and Recombinant DNA Technology

Types of Vectors

Cloning vectors provide a backbone for the DNA insert to be reproduced and propagated in bacteria however, these vectors are only useful for storing a genetic sequence. By themselves, they are incapable of allowing for transcription and translation of the gene into a functional protein product.

For a gene to give rise to a protein product, an expression vector must be used that contains the necessary elements for a host cell to transcribe and translate the gene. In the case of a mammalian cell, a standard mammalian expression vector will contain an origin of replication, MCS, and selectable marker, as described above. However, the expression vector will also need a promoter found in mammalian cells that can drive the expression of the gene. The coding DNA needs other features to be transcribed and translated, such as the polyadenylation tail that normally appears at the end of transcribed pre-mRNA and a sequence that attracts the ribosome for translation.

Key Terms

  • plasmid: A circle of double-stranded DNA that is separate from the chromosomes, which is found in bacteria and protozoa.
  • expression vector: An expression vector, otherwise known as an expression construct, is generally a plasmid that is used to introduce a specific gene into a target cell.
  • transcription: The synthesis of RNA under the direction of DNA.

An expression vector, otherwise known as an expression construct, is generally a plasmid that is used to introduce a specific gene into a target cell. Once the expression vector is inside the cell, the protein that is encoded by the gene is produced by the cellular-transcription and translation machinery ribosomal complexes. The plasmid is frequently engineered to contain regulatory sequences that act as enhancer and promoter regions and lead to efficient transcription of the gene carried on the expression vector. The goal of a well-designed expression vector is the production of large amounts of stable messenger RNA, and in extension, proteins. Expression vectors are basic tools for biotechnology and the production of proteins such as insulin, which is important for the treatment of diabetes.

Figure: The pGEX-3x Plasmid: The pGEX-3x plasmid is a popular cloning vector. Please note the presence of a multiple cloning site, a promoter, a repressor, and a selectable marker.

After expression of the gene product, the purification of the protein is required but since the vector is introduced to a host cell, the protein of interest should be purified from the proteins of the host cell. Therefore, to make the purification process easy, the cloned gene should have a tag. This tag could be histidine (His) tag or any other marker peptide.

Expression vectors are used for molecular biology techniques such as site-directed mutagenesis. Cloning vectors, which are very similar to expression vectors, involve the same process of introducing a new gene into a plasmid, but the plasmid is then added into bacteria for replication purposes. In general, DNA vectors that are used in many molecular-biology gene-cloning experiments need not result in the expression of a protein.

Expression vectors must have expression signals such as a strong promoter, a strong termination codon, adjustment of the distance between the promoter and the cloned gene, and the insertion of a transcription termination sequence and a PTIS (portable translation initiation sequence).

A shuttle vector is a vector that can propagate in two different host species, hence, inserted DNA can be tested or manipulated in two different cell types. The main advantage of these vectors is that they can be manipulated in E. coli and then used in a system which is more difficult or slower to use.

Shuttle vectors can be used in both eukaryotes and prokaryotes. Shuttle vectors are frequently used to quickly make multiple copies of the gene in E. coli (amplification). They can also be used for in vitro experiments and modifications such as mutagenesis and PCR. One of the most common types of shuttle vectors is the yeast shuttle vector that contains components allowing for the replication and selection in both E. coli cells and yeast cells. The E. coli component of a yeast shuttle vector includes an origin of replication and a selectable marker, such as an antibiotic resistance like beta lactamase. The yeast component of a yeast shuttle vector includes an autonomously replicating sequence (ARS), a yeast centromere (CEN), and a yeast selectable marker.

Genetically engineered plasmid can be used to fight antimicrobial resistance

Researchers have engineered a plasmid to remove an antibiotic resistance gene from the Enterococcus faecalis bacterium, an accomplishment that could lead to new methods for combating antibiotic resistance. The research is published this week in Antimicrobial Agents and Chemotherapy, a journal of the American Society for Microbiology.

In vitro, and in mouse models, the engineered plasmid removed the antibiotic resistance gene from E. faecalis. In mouse models, it reduced the abundance of the resistance gene threefold..

"Our concern with organisms that cause hospital-acquired infections that are resistant to many clinical antibiotic therapies motivated the research," said co-senior author Breck A. Duerkop, PhD, Assistant Professor of Immunology and Microbiology, University of Colorado School of Medicine, Anschutz Medical Center, Aurora.

E. faecalis is part of the normal, benign intestinal flora, but when antibiotics kill off beneficial intestinal flora, E. faecalis can become pathogenic. As such, it can also acquire single or multidrug resistance. Antibiotic resistant E. faecalis infections are a major problem in hospitals.

The mechanism used to remove antibiotic resistance genes is the specialized protein, CRISPR-Cas9. It can make cuts just about anywhere in DNA.

Along with CRISPR-Cas9, RNA sequences homologous to DNA within the antibiotic resistance gene have been added to the engineered plasmid. These RNAs guide the CRISPR-Cas9 to make the cuts in the right places.

Previous work in animal models by co-senior investigator Kelli L. Palmer, PhD, found that CRISPR-Cas9 could prevent intestinal E. faecalis from acquiring resistance genes. Dr. Palmer is Fellow, Cecil H. and Ida Green Chair in Systems Biology Science, Associate Professor of Biological Sciences, University of Texas, Dallas.

The delivery vehicle for the engineered plasmid is a particular strain of E. faecalis, which conjugates with E. faecalis of various different strains. Conjugation is the process whereby bacteria come together to transfer genetic material from one to the other via direct cell to cell contact.

"E. faecalis strains used to deliver these plasmids to drug resistant strains [of E. faecalis] are immune to acquiring drug resistant traits carried by the target cells," said Dr. Duerkop. "The engineered plasmid can significantly reduce the occurrence of antibiotic resistance in the target bacterial population rendering it more susceptible to antibiotics. We envision that this type of system could be used to re-sensitize antibiotic resistant E. faecalis to antibiotics," he said.

Nonetheless, Dr. Duerkop cautioned that it remained possible that E. faecalis could still circumvent the engineered plasmid. Some bacteria have anti-CRISPR systems that can block CRISPR-Cas9 function, and some others have systems that can degrade foreign DNA. "Future studies will need to be done to address such an issue as E. faecalis avoiding the targeting system and under what conditions this may happen," said Dr. Duerkop.

Designing plasmid vectors

Nonviral gene therapy vectors are commonly based on recombinant bacterial plasmids or their derivatives. The plasmids are propagated in bacteria, so, in addition to their therapeutic cargo, they necessarily contain a bacterial replication origin and a selection marker, usually a gene conferring antibiotic resistance. Structural and maintenance plasmid stability in bacteria is required for the plasmid DNA production and can be achieved by carefully choosing a combination of the therapeutic DNA sequences, replication origin, selection marker, and bacterial strain. The use of appropriate promoters, other regulatory elements, and mammalian maintenance devices ensures that the therapeutic gene or genes are adequately expressed in target human cells. Optimal immune response to the plasmid vectors can be modulated via inclusion or exclusion of DNA sequences containing immunostimulatory CpG sequence motifs. DNA fragments facilitating construction of plasmid vectors should also be considered for inclusion in the design of plasmid vectors. Techniques relying on site-specific or homologous recombination are preferred for construction of large plasmids (>15 kb), while digestion of DNA by restriction enzymes with subsequent ligation of the resulting DNA fragments continues to be the mainstream approach for generation of small- and medium-size plasmids. Rapid selection of a desired recombinant plasmid against a background of other plasmids continues to be a challenge. In this chapter, the emphasis is placed on efficient and flexible versions of DNA cloning protocols using selection of recombinant plasmids by restriction endonucleases directly in the ligation mixture.

Genetic engineering in plants

Because of their economic significance, plants have long been the subject of genetic analysis aimed at developing improved varieties. Recombinant DNA technology has introduced a new dimension to this effort because the genome modifications made possible by this technology are almost limitless. No longer is breeding confined to selecting variants within a given species. DNA can now be introduced from other species of plants, animals, or even bacteria.

Ti plasmid.  

The only vectors routinely used to produce transgenic plants are derived from a soil bacterium called Agrobacterium tumefaciens. This bacterium causes what is known as crown gall disease, in which the infected plant produces uncontrolled growths (tumors, or galls), normally at the base (crown) of the plant. The key to tumor production is a large (200-kb) circular DNA plasmid — the Ti (tumor- inducting) plasmid. When the bacterium infects a plant cell, a part of the Ti plasmid — a region called T-DNA — is transferred and inserted, apparently more or less at random, into the genome of the host plant (Figure 13-14). The functions required for this transfer are outside the T-DNA on the Ti plasmid. The T-DNA itself carries several interesting functions, including the production of the tumor and the synthesis of compounds called opines. Opines are actually synthesized to the host plant under the direction of the T-DNA. The bacterium then uses the opines for its own purposes, calling on opine-utilizing genes on the Ti plasmid. Two important opines are nopaline and octopine two separate Ti plasmids produce them. The structure of Ti is shown in Figure 13-15.

Figure 13-14

In the process of causing crown gall disease, the bacterium A. tumefaciens inserts a part of its Ti plasmid — a region called T-DNA — into a chromosome of the host plant.

Figure 13-15

Simplified representation of the major regions of the Ti plasmid of A. tumefaciens. The T-DNA, when inserted into the chromosomal DNA of the host plant, directs the synthesis of nopaline, which is then utilized by the bacterium for its own purposes. T-DNA (more. )

The natural behavior of the Ti plasmid makes it well suited to the role of a plant vector. If the DNA of interest could be spliced into the T-DNA, then the whole package would be inserted in a stable state into a plant chromosome. This system has indeed been made to work essentially in this way, but with some necessary modifications. Let’s examine one protocol.

Ti plasmids are too large to be easily manipulated and cannot be readily made smaller, because they contain few unique restriction sites. Consequently, a smaller, intermediate vector initially receives the insert of interest and the various other genes and segments necessary for recombination, replication, and antibiotic resistance. When engineered with the desired gene elements, this intermediate vector can then be inserted into the Ti plasmid, forming a cointegrate plasmid that can be introduced into a plant cell by transformation. Figure 13-16a shows one method of creating the cointegrate. The Ti plasmid that will receive the intermediate vector is first attenuated that is, it has the entire right-hand region of its T-DNA, including tumor genes and nopaline-synthesis genes, deleted, rendering it incapable of tumor formation — a “nuisance” aspect of the T-DNA function. It retains the left-hand border of its T-DNA, which will be used as the crossover site for incorporation of the intermediate vector. The intermediate vector has had a convenient cloning segment spliced in, containing a variety of unique restriction sites. The gene of interest has been inserted at this site in Figure 13-16. Also spliced into the vector are a selectable bacterial gene (spc R ) for spectinomycin resistance a bacterial kanamycin-resistance gene (kan R ), engineered for expression in plants and two segments of T-DNA. One segment carries the nopaline-synthesis gene (nos) plus the right-hand T-DNA border sequence. The second T-DNA segment comes from near the left-hand border and provides a section for recombination with a homologous part of the left-hand region, which was retained in the disarmed Ti plasmid. After the intermediate vectors have been introduced into Agrobacterium cells containing the disarmed Ti plasmids (by conjugation with E. coli), plasmid recombinants (cointegrates) can be selected by plating on spectinomycin. The selected bacterial colonies will contain only the Ti plasmid, because the intermediate vector is incapable of replication in Agrobacterium.

Figure 13-16

(a) To produce transgenic plants, an intermediate vector of manageable size is used to clone the segment of interest. In the method shown here, the intermediate vector is then recombined with an attenuated (𠇍isarmed”) Ti plasmid to generate (more. )

As Figure 13-16b shows, after spectinomycin selection for the cointegrates, bacteria containing the recombinant double, or cointegrant, plasmid are then used to infect cut segments of plant tissue, such as punched-out leaf disks. If bacterial infection of plant cells takes place, any genetic material between the left and right T-DNA border sequences can be inserted into the plant chromosomes. If the leaf disks are placed on a medium containing kanamycin, the only plant cells that will undergo cell division are those that have acquired the kan R gene from T-DNA transfer. The growth of such cells results in a clump, or callus, which is an indication that transformation has taken place. These calli can be induced to form shoots and roots, at which time they are transferred to soil where they develop into transgenic plants (Figure 13-16b). Often only one T-DNA insert is detectable in such plants, where it segregates at meiosis like a regular Mendelian allele (Figure 13-17). The insert can be detected by a T-DNA probe in a Southern hybridization or can be verified by the detection of the chemical nopaline in the transgenic tissue.

Figure 13-17

T-DNA and any DNA contained within it are inserted into a plant chromosome in the transgenic plant and then transmitted in a Mendelian pattern of inheritance.

Expression of cloned DNA

The DNA cloned into the T-DNA can be any DNA that the investigator wants to insert into the subject plant. A particularly striking foreign DNA that has been inserted with the use of T-DNA is the gene for the enzyme luciferase, which is isolated from fireflies. The enzyme catalyzes the reaction of a chemical called luciferin with ATP in this process, light is emitted, which explains why fireflies glow in the dark. A transgenic tobacco plant expressing the luciferase gene also will glow in the dark when watered with a solution of luciferin (see photograph at the beginning of this chapter). Such manipulation might seem like an attempt to develop technology for making Christmas trees that do not need lights, but in fact the luciferase gene is useful as a reporter to monitor the function of any gene during development. In other words, the upstream promoter sequences of any gene of interest can be fused to the luciferase gene and put into a plant by T-DNA. Then the luciferase gene will follow the same developmental pattern as that of the normally regulated gene, but the luciferase gene will announce its activity prominently by glowing at various times or in various tissues, depending on the regulatory sequence.

Other genes used as reporters in plants are the bacterial GUS (β-glucuronidase) gene, which turns the compound X-Gluc blue, and the bacterial lac (β-galactosidase) gene, which turns X-Gal blue. Cells in which these reporters are expressed turn blue, and this blueness can be easily seen either by the naked eye or under the microscope.

Transgenic plants carrying any one of a variety of foreign genes are in current use, and many more are in development. Not only are the qualities of plants themselves being manipulated, but, like microorganisms, plants are also being used as convenient �tories” to produce proteins encoded by foreign genes.

Why would a mammalian vector plasmid require an antibacterial resistance gene? - Biology

A major thrust of vector development has been to create vectors that will handle larger foreign DNA inserts, aiming to reduce the number of recombinants it is necessary to look at in order to identify a specific DNA sequence.

Cosmid vectors were among the first large insert cloning vehicles developed.

The vector replicates as a plasmid
(it contains a ColE1 origin of replication),
uses Amp r for positive selection and
employs lambda phage packaging to select for recombinant plasmids carrying foreign DNA inserts 45 KB in size.

Ligation of cosmid vector and foreign DNA fragments (SauIIIA partial digest fragments 45 kb in size) is similar to ligation into a lambda substitution vector.
The desired ligation product is a concatemer of
45 kb foreign DNA fragment and 5 kb cosmid vector sequences.

This concatemer is then packaged into viral particles (remember packaging is cos site to cos site) and these are used to infect E. coli where the cosmid vector replicates using the ColE1 orignin of replication. Phage packaging serves only to select for recombinant molecules and to transfer these long DNA molecues (50 kb total) into the bacterial host (50 kb fragments transform very inefficiently while phage infection is very efficient).

Recently, the need for prokaryotic vectors capable of replicating very large foreign DNA fragments
(> 1 MB in size) has produced a couple of additional prokaryotic vector systems.
One is based on another bacteriophage called P1.
P1 phage can carry in excess of 100 KB of foreign DNA but do not seem to have found wide acceptance.

A few years ago, a Bacterial Artificial Chromosome (BAC) vector was developed allowing foreign DNA fragements over 1 MB to be propagated in E. coli.
These BAC vectors replicate as low copy number plasmids (to minimize the demands on the host replication system) but how do we get such large DNA fragments into E. coli?

Standard methods for transforming E coli rely on the chemical preparation of the E coli to put them in a 'transformation competent' state.
This typically involves incubating the bacteria in the presence of divalent cations (particularly Ca +2 ) at low temperature (4 o C) which puts the cell membrane into a 'semi-crystaline state. Addition of DNA to these 'competent cells' allow the DNA to bind to the semi-crystaline membrane. Heat shocking the cells (42 o C) remobilizes the membrane and allows the DNA to cross the membrane. This transit across the membrane is sensitive to DNA length - small plasmids cross the membrane rapidly while larger molecules are unable to cross.

Getting very large molecules across the cell membrane relies on an alternative mechanism called electroporation. This involves mixing DNA with E coli cells and then exposing the mixture to a high-voltage pulse. The high voltage induces massive membrane rearrangements and the coincident uptake of DNA which is independant of size. BACs have been an important tool for the human genome sequencing project.

Numerous vector systems have been developed for use in eukaryotic systems.
The most common of these are often called 'shuttle vectors' as they replicate in both prokaryotic and eukaryotic hosts. In general, DNA manipulations and characterizations are done in prokaryotic systems, and then the manipulated DNA is reintroduced to eukaryotic systems for functional analysis.

Yeast Vectors
Shuttle vectors were first developed for transfering DNA fragments between E coli and S cerevisiae. These vectors contain an antibiotic resistance gene for positive selection in E coli, and E coli origin of replication (ColE1 ori), and a polylinker in the lacZ alpha-complementing fragment for insertional inactivation by the foreign DNA insert. In addition, the vector contains a yeast specific origin of replication (derived from the naturally occuring yeast plasmid called the 2 um circle - 2um ori)
and amino-acid or N-base biosynthsis enzymes for positive selection in yeast (HIS, LEU and ADE genes)

Mammalian Vectors
Shuttle vectors have also been developed for use in mammalian tissue culture.
Like the yeast shuttle vectors, mammalian shuttle vectors require sequeces enabling them to replicate in mammalian tissue culture. These eukaryotic origins of replication are typically derived from well characterized mammalian viruses - most commonly SV-40 (SV-40 ori and Large T antigen system) or Epstein-Barr virus (mononucleosis). Both systems allow the vector to replicate within the host cell as a plasmid without integrating into the genome. In addition to the origin of replication, these shuttle vectors also carry antibiotic resistance genes which function in eukaryotic cells (ie neomycin (G418) resistance, hygromycin resistance, methotrexate resistance etc).

Finally, artificial chromosome vectors have been developed for use in yeast (YACs) and are being developed for use in mammalian cells (MACs).
These eukaryotic vectors contain telomeric sequences (eukaryotic chromsomes are linear not circular molecules) and centromeres to ensure appropriate segregation of the artificial chromosomes as well as selectable markers. Such vector systems may gain in importance as we move towards manipulating the eukaryotic genome.


Heterologous multigene expression in mammalian cells is a key technology in contemporary biological research. Unfortunately, all available technologies suffer from major deficiencies such as unreliable co-transfection efficiency. Our MultiLabel system is clearly superior to the currently available technology for a number of reasons. Several expression cassettes can be assembled in a combinatorial way because the system is modular. This is a clear advantage when combinations of proteins or mutations of a protein need to be tested. All cells express the entire set of transfected genes. Therefore, the population of transfected cells is uniform and not a mixture of cells expressing only a fraction of the transfected genes, a fact that is especially important in biochemical assays. Simultaneous insertion of several genes into a cell genome is possible in a single step, which is less time-consuming than sequential integration by multiple transfections. The expression levels can be regulated independently because the system transcribes autonomous mRNAs and is not polycistronic. The use of IRES elements often leads to unpredictable expression of the adjacent genes. Several cloning strategies, including robotics-based high-throughput methods, can be used because of the application of standardized cloning sites. The assembly of the expression cassettes is recombination based and therefore independent of restriction enzymes. Our design also allows the rapid modification of acceptor and donor vectors, thus extending the scope of our technology to viral expression or to different integration systems.

Our results provide evidence for the first time that it is possible to use this technology concept to perform cell biological assays with high efficacy. Our results clearly demonstrate that the behaviour of the transfected cells is not negatively affected, that we can reproduce previously published intracellular trafficking studies and that the transfected cells are evidently still able to proliferate. Most contemporary time-lapse movies and time-course studies typically show only two colour channels. In contrast, we show here a time course of a transiently transfected cell with four channels and a movie of a cell line with five channels. Taken together, this provides compelling evidence that MultiLabel indeed addresses and alleviates hitherto inhibiting technical problems.

We anticipate that a broad range of applications in cell biology will significantly benefit from the MultiLabel system. In this study, using fluorescent markers, we demonstrate the superior performance of MultiLabel when compared with conventional coexpression systems. By introducing affinity purification tags, our approach will be of equal benefit for systems and structural biology applications in mammalian cells. We considerably extend the existing tool box for expression of protein and protein complexes, for instance, in the context of applications in drug development and screening.

AAV VECTOR MANUFACTURING – Challenges & Opportunities in the Manufacturing of AAV Vectors Used in the Delivery of Gene Therapy Treatments

Significant advances in the specific targeting of delivery vectors and the increased therapeutic efficacy of such vectors for gene delivery have been made, stimulating major interest in the development and commercialization of therapeutic products focused on gene therapy indications. In the past few years, there have been a large number of positive clinical outputs for gene therapy-based products spanning broad therapeutic areas, including CAR T-cell immunotherapy, oncology, and regenerative medicine based on monogenetic diseases. In terms of gene delivery, viral vectors have emerged as the preferred vehicles of choice, used in 48% of the 483 current on-going gene therapy trials. 1

Of the gene therapy products in development, recombinant Adeno-Associated Virus (AAV)-based vectors are currently the most widely used and show the greatest potential for delivery in gene therapy indications. 1-3 The first rAAV-vector-based clinical trial was performed 20 years ago a Phase I study delivering a CTFR transgene via an Raav vector to adult cystic fibrosis patients with mid lung disease. 4 In 2015, there were 103 rAAV vector-based products reported to be in development, a number that is expected to increase further in the next few years. 1 The preferred use of rAAV vector systems is due, in part, to the lack of disease associated with the wild-type virus, the ability of AAV to transduce non-dividing as well as dividing cells, and the resulting long-term robust transgene expression observed in several Phase I/II trials. 5 Furthermore, different rAAV vector serotypes, either naturally occurring or hybrid/synthetically derived, can be exploited to specifically target different tissues, organs, and cells, and help evade any pre-existing immunity to the vector, thus expanding the therapeutic application and commercial potential of AAV-based gene therapies. 6

Whilst the majority (74%) of AAV-based therapeutic products are in early to mid-phase clinical development, a number of promising clinical outputs have helped progress the pipeline of possible AAV-based products. In 2012, the Dutch-based company UniQure was granted marketing authorization in Europe for Glybera ® , an AAV1-based gene therapy for the treatment of adult patients diagnosed with familial lipoprotein lipase deficiency (LPLD). The recent clinical successes of AAV-, and other viral vector-based gene therapies has fueled significant investments into the sector as such, there is growing demand for clinical-grade good manufacturing practice (GMP) production solutions for these viral vector products.


The development of manufacturing processes for novel biotherapeutics is time consuming and expensive. Recombinant viral vector production is seen as complex, with the production scale-up regarded as a major challenge technically, and a large barrier for commercialization.

Reported clinical doses for AAV-based viral vectors range from 1011 to 1014 genomic particles (vector genomes vg) per patient dependent on therapeutic area. 1,3,6 From a wider gene therapy development perspective, current scale-out approaches fall short of supplying the required number of doses to allow later Phase (II/III) trails to progress, thus retarding the development of gene therapy products. This is supported by the fact that the majority of clinical studies have been very small, performed on <100 patients (and in some cases <10), using adherent cell transfection processes that generate very modest amounts of product. When predicted amounts of virus required for later phase development are compared to current productivities (ca. 5x 1011 vg from single 10 layer cell factory), there is real concern that this approach will fall short of the material requirements for late phase and in-market needs for even ultra-orphan diseases, which have high dose and small patient cohorts, let alone more “standard” gene therapy indications.

In the case of AAV, a number of production strategies exist to generate viral vectors as with all different strategies a number of advantages and disadvantages are associated with each.


Generation of stable engineered cell lines, through the introduction of both regulatory (Rep) and structural capsid (Cap) genes and/or the rAAV genome, give rise to packaging or producer cell lines, respectively. AAV viral vectors are produced from packaging cell lines following transfection of the AAV construct and the co-infection with a helper virus, such as adenovirus (Ad) or Herpes Simplex Virus (HSV) or via a single infection with a recombinant helper viral vector containing the rAAV genome. For producer cell lines, AAV is generated following a single-step infection with an Ad or HSV helper virus. Stable cell lines have been reported to produce relatively high AAV vector genome (vg) particles per cell (up to 10,000 vg per producing cell). Packaging and producer cell lines have been generated using cell lines capable of both adherent and suspension growth, allowing for processes to be developed that utilize traditional tissue culture systems for small scale, combined with larger-scale manufacturing performed in suspension bioreactors. This scalable approach has been used in the production of an AAV1 product for heart failure here the viral vector has been generated at a 2000-L scale from a HeLaS3 producer cell line, following coinfection with helper viruses. 7

Despite the relatively high yield and scalability reported, a number of disadvantages have limited the use of the producer and packaging cell lines in clinical development. The generation of stable lines is technically demanding, extremely time consuming, and needs to be performed for every vector and AAV serotype combination. Clonal characterization and cell line stability is very time consuming and expensive, and furthermore carries potential risk based around cell passage history and the relationship between growth kinetics and vector production. A major concern is the use of helper Ad and hybrid variants in these systems. The production of the helper virus can itself be extremely costly and lengthy, with the need to carefully assess the quality of this critical starting material ahead of use in AAV production. Furthermore, establishing effective removal and clearance procedures to separate the helper virus away from the AAV vector product is not trivial, resulting in complex and costly downstream process development and a potentially high cost of goods.


Production of AAV vectors using a Baculovirus (BV) expression system emerged as a consequence of the BV system’s ability to produce complex glycosylated recombinant proteins at high levels in SF9 insect cells at high cell densities. Building on AAV production via a stable cell line system, the BV system was developed to produce viral vector without the need to co-infect with a human helper virus. AAV2 was produced in Sf9 insect cells following co-infection with three recombinant BV vectors, encoding the transgenes for Rep, Cap, and rAAV genome, respectively. Since then, the BV system has been modified and improved, with systems now developed that use a two-vector approach, reducing the complexity of the process. Scalable solutions have also been employed one approach used cryopreserved BV-infected insect cells that separately carry the required AAV components (rAAV genome, Rep, and Cap genes). The infected cell were used to inoculate a 200-L scale bioreactor containing unmodified insect cells, and released infectious rBV (rAAV, rCAP, rRep) in a continuous manner that subsequently infected the uninfected cells, thus driving a sustained production phase. 8

Currently, the AAV1-based drug Glybera is produced using the BV system however, several drawbacks have limited the use of the BV system to produce AAV vectors in the clinical setting. Overcoming the molecular cell biology aspects needed to produce a mammalian viral vector in an insect cell system using insect virus has major challenges instability of AAV genes within BV has been reported, along with the inability to assemble AAV particles with the correct stoichiometry of capsid proteins, affecting the infectivity of the produced AAV viral vector. Similar to the stable cell line production systems, there are challenges in clearing and removing the starting and propagated BV from the AAV viral vectors during the downstream purification operation. However, several biopharmaceutical companies are currently using variations of the BV platform, and are actively working on improvements to mitigate the limitations of the BV system, moving toward large-scale clinical rAAV manufacture.


Transient transfection of plasmid DNA into mammalian cells for the production of AAV viral vectors is the strategy most commonly used in clinical grade manufacturing of these viral vectors. rAAV vectors are usually produced in human embryonic kidney 293 cells (HEK293), or HEK293 cell variants following introduction (transfection) of typically three DNA plasmids carrying the regulatory (Rep) and structural capsid (Cap) genes, the rAAV transgene, and the specific genes that provide helper Ad function. If all three plasmids are successfully transfected into a cell, the cell will produce an rAAV vector without the need to co-infect with wild type helper virus. The transfection approach is fairly rapid and versatile and has been used to produce different serotypes of rAAV, as only the gene sequence of the Cap genes has to be altered to produce the various serotypes. Furthermore, modification of the transgene plasmid allows for both single- and double-stranded (self-complementary) forms of the vector to be generated.

Again, the main challenge to this approach is the inherent lack of scalability due to the use of adherent HEK293 cells. As such, adherent cell systems require a scale-out approach based on the linear expansion of 2D surface area, rather than the volumetric 3D scale-up approach usually employed in the production of biopharmaceuticals. In order to produce and purify the required vector genome (vg) particle numbers of >1� vg to support mid- to later-stage clinical trials, over 500 Hyperstacks™ (a Hyperstack has 36 layers and total surface area of 18,000 cm 2 ) would be needed. Whilst possible, this approach is not a viable option for most manufacturing facilities due to facility footprint, cost, and man power limitations. Work is underway to develop truly scalable, regulatory compliant, production solutions for rAAV vector generation. HEK293 cells have been adapted to suspension growth in animal component-, serum-free, and antibiotic-free media systems, with optimization of transfection conditions evaluated in shake flask, rocking, and stirred-tank bioreactor systems.


Irrespective of vector production processes and scales, the ultimate goal of for AAV vector manufacturing is to have robust downstream purification that generates final clinical material that has high titre, high potency, and high purity. Due to the emerging nature of the viral vector field, most downstream approaches are based around traditional laboratory processes that are not scalable or suitable for clinical-grade manufacture. Limitations exist throughout the downstream purification of viral vectors these include: the harvesting of the producer cells, cell lysis procedures to release AAV vector, the clarification and removal of cellular impurities, vector separation and purification, and vector formulation and sterile filtration.

The challenge can be exemplified by the way that genomic-containing vector particles have been traditionally separated from empty particles using gradient ultra-centrifugation. Whilst the resulting material obtained from such a step is of high purity and high potency, the time-consuming nature and complexity of scale up significantly limits the use of ultra-centrifugation within the downstream process. For therapeutic indications, other than those focused at ultra-orphan diseases that require small volumes of material to be purified, the need for processing large volumes requires an alternative method of purification. As such, chromatography-based approaches are being developed using affinity binding and/or ion-exchange steps that provide efficiency, flexibility, and scalability to the purification of AAV vectors. As high-purity, high-potency vectors can be generated using ultracentrifugation approaches, the challenge for a chromatographic purification approach is to generate vectors with the same degree of purity and potency. Regulatory consideration must therefore be given to any process change during the clinical development program. Also, due to the chemical and biological variances observed between the various AAV serotypes, the development of a single downstream platform for all AAV vectors may be unlikely rather a number of potential solutions are likely to be employed.


Developments are underway to improve clinical manufacturing processes and to move into scalable and controllable production and purification systems. However, these are stymied by the lack of processing knowledge of critical parameters at all stages of the manufacturing process, driven by the lack of suitable analytical systems to support development studies, and to interrogate and control production processes, as well as characterize final materials. 9 Linked to the manufacturing challenge is the regulatory requirement for the identification, characterization, and batch-to-batch control of the process- and product-related impurities present in highly purified material. Especially challenging are impurities related to the AAV vector products, which closely resemble the vector itself.

AAV vector particles can co-package non-specific plasmid material into the capsid. It has been reported that between 1% and 8% of purified AAV particles contain an incorrect nucleic acid sequence. 7 As reported clinical dose ranges vary from 1011 to 1014 genomic particles, there could be as many as 109 co-packaged impurities per dose. These process-related impurities require detailed characterization, along with an evaluation of the potential for the incorrectly packaged DNA to be active in transduced cells. 10

Interestingly, most plasmids currently used for transient transfection carry an antibiotic resistance gene to enable plasmid selection and maintenance during their own production. Therefore, removal of antibiotic resistance genes would be desirable for the production of AAV in Helper-free systems. Technology exists, such as Cobra’s patented Operator Repressor Titration (ORT™) plasmid maintenance system, to generate plasmid DNA lacking antibiotic resistance genes. 11 These systems eliminate the chance of co-packaging a functional antibiotic-resistance gene into the viral capsid, thus increasing the safety of the product and reducing the burden of characterization of AAV-related impurities.


There remains a clear need for improved process productivities, and the need to develop manufacturing processes that can be applied to a wide number of AAV-based viral vector therapeutic candidates. Simplistically, the AAV vector is a delivery vehicle for a therapeutic gene, and the manufacturing process is not linked to that gene. Logic therefore dictates that it should be possible to generate platform processes specific for AAV serotypes and even possible to generate processes that can be applied to multiple serotypes. This approach is not unprecedented the development of platform-based development and manufacturing processes that disconnects production from specific products has had an enormous impact in other biologic areas (ie, monoclonal antibodies), reducing development cost, timelines, and cost of goods, and has also allowed drug developers to significantly expand clinical pipelines in multiple therapeutic areas.

To achieve this for AAV vectors, the underpinning scientific knowledge surrounding critical operational parameters in the manufacturing process needs to be generated and optimized, along with a suite of standard analytical methods. Any platform developed will need to be sufficiently flexible to accommodate multiple AAV vector types. Such development will have long-term benefits to the generation of materials for first-in-man studies, and onward for late-stage clinical studies and ultimately in-market supply.

1. Gene Therapy Market 2015-2025, Roots Analysis Report (2015).
2. Ginn SL, Alexander IE, Edelstein ML, Abedi MR, Wixon J. Gene therapy clinical trials worldwide to 2012 – an update. J Gene Med. 201315:65-77.
3. Okada. Gene Therapy, Tools & Potential Application. 2013C17.
4. Flotte, et al. Hum Gene Ther. 19967:1145-1159.
5. Okada, et al. Methods Enzymol. 2002346:378-393.
6. Wu, et al. Molecular Therapy. 200614:316-327.
7. Hajjar et al. J Card Fail. 200814:355-367.
8. Cecchini et al. Hum Gene Ther 201122:1021-1030.
9. Wright JF. Biomedicines. 20142:80-97.
10. Quality, non-clinical and clinical issues related to the development of rec. AAV vectors 24 June 2010,EMEA/CHMP/GTWP/587488/2007.
11. Cranenburgh. Et al. Nucleic Acids Res. 200129:e26.

To view this issue and all back issues online, please visit .

Dr. Daniel C. Smith is Chief Scientific Officer of Cobra Biologics with the responsibility for developing the highest level of scientific excellence across the Group and enhancing Cobra’s DNA, virus, microbial, and mammalian proteins research and development platforms. Prior to joining Cobra, he spent 4 years with the bioProcessUK team at the HealthTech & Medicines Knowledge Transfer Network (KTN), driving the innovation agenda for biologics bioprocessing in the UK as a Knowledge Transfer Manager. Dr. Smith gained his industrial experience at Cobra in a variety of roles progressing from Senior Scientist to Commercial Scientific Development Manager, responsible for developing the strategy for customer’s projects, alongside maintaining scientific oversight of Cobra’s R&D projects. He earned his BSc (Hons) in Biochemistry and his PhD in Molecular Cell Biology. He has more than 20 research publications to his credit has worked with a number of academic groups across the UK, Europe, and the US and has extensive academic research experience in cell biology, molecular biology, immunology, and protein biochemistry, with a particular expertise in protein-based toxins and cellular delivery vectors.

Transferring the Gene

A vector is a carrier DNA molecule into which the desired gene can be inserted.

Most commonly, this vector is a plasmid. This is a small, extra-chromosomal, circular piece of DNA often found in bacteria in addition to their functional DNA.

The plasmids are modified so that they have two or more genes for resistance to antibiotics. They should also contain a sequence that can be recognised by the same restriction enzyme used to cut the fragments. The site that is cut should be in one of the genes for antibiotic resistance.

A Plasmid cut by Bam HI:

How the gene is introduced into the plasmid:

Cut the genome with a restriction enzyme (RE) and mix with the plasmid that has also been cut with the same R.E so that the sticky ends of the fragments and the plasmid are complementary. Hopefully, some fragments will insert into the plasmid DNA before either segment joins with itself.

The fragments are added to the plasmids with these possible outcomes:

  1. Plasmid rejoins, tetracycline gene now intact.
  2. Fragment joins with plasmid. Tetracycline resistance gene is interrupted the fragment does not contain the desired gene.
  3. Fragment joins with plasmid. Tetracycline gene is interrupted this time the fragment does contain the desired gene.
  4. The fragment joins with itself.

Some plasmids will now contain the recombinant DNA fragment.

Other plasmids, however will not contain a fragment. If the plasmids are recombinants then one of the antibiotic resistance genes (e.g, for tetracycline) will have been disrupted. However, the other gene for antibiotic resistance (e.g. for ampicillin) will still be intact.

There is still one problem - how do you identify the recombinant plasmid with the desired gene in it?

Isolating the desired gene in the right colony

You need to know the DNA base sequence of the gene for the desired protein so that a section of that base sequence can be radioactively labelled.

This section of DNA with the correct base sequence is called a probe.

The DNA in the bacteria is "unzipped" so that it becomes single stranded and a probe would anneal (attach) if there were complementary bases.

The probe is added and sticks to the correct complementary fragment. The correct fragment can now be identified, as it is radioactive.

All bacteria in a colony have been produced by the replication of one individual. They will all therefore have the same genes to produce the same proteins. This colony can then be isolated and multiplied so that the protein is synthesised and can be harvested for further use.

Bacteria can be used in this way to produce human insulin for diabetics who do not produce their own.

PCOR: a new design of plasmid vectors for nonviral gene therapy

A totally redesigned host/vector system with improved properties in terms of safety has been developed. The pCOR plasmids are narrow-host range plasmid vectors for nonviral gene therapy. These plasmids contain a conditional origin of replication and must be propagated in a specifically engineered E. coli host strain, greatly reducing the potential for propagation in the environment or in treated patients. The pCOR backbone has several features that increase safety in terms of dissemination and selection: (1) the origin of replication requires a plasmid-specific initiator protein, π protein, encoded by the pir gene limiting its host range to bacterial strains that produce this trans-acting protein (2) the plasmid’s selectable marker is not an antibiotic resistance gene but a gene encoding a bacterial suppressor tRNA. Optimized E. coli hosts supporting pCOR replication and selection were constructed. High yields of supercoiled pCOR monomers were obtained (100 mg/l) through fed-batch fermentation. pCOR vectors carrying the luciferase reporter gene gave high levels of luciferase activity when injected into murine skeletal muscle.

Two different types of DNA vehicles, based on recombinant viruses and bacterial DNA plasmids, are used in gene therapy. Although virus-based vectors are widely used and very efficient vectors, the potential risks of viral gene delivery have to be seriously considered. E. coli plasmid-based vectors are non-infectious and versatile vehicles for gene therapy. The manufacture of therapeutic plasmid DNA that carries potent human genes becomes a reality. Thus, new approaches that take into account safety and environmental consideration must be developed.

Vector design is a major contributor to the safety of production and therapy. Efforts to improve the plasmid backbone have focused on the improvement of transgene expression1 or plasmid copy number2 but no rational design in terms of dissemination or size, has been reported. The ColE1-derived bacterial plasmids are currently used in gene therapy experiments and clinical trials. As far as the safety and containment of such genetic material are concerned, two elements are deleterious: the ColE1 origin of replication and the antibiotic resistance marker. Clinical use of such plasmids could potentially lead to their dissemination in the environment or in the patient. This could be of the utmost importance for the immunodepressed or for cystic fibrosis patients who suffer from chronic bronchopulmonary infections.3 pCOR plasmids, for plasmid with conditional origin of replication, were specifically designed and developed for gene therapy. These new plasmids are small efficient transgene carriers and fulfill several quality assurance and safety criteria.4 These high copy number plasmids carry a minimal amount of bacterial sequences, a conditional origin of replication that prevents dissemination during production or therapy and do not contain an antibiotic resistance gene.

The pCOR backbone consists of three bacterial elements: the 0.4 kb R6K γ conditional origin of replication (ori γ), which requires the R6K π initiator protein to be functional, a 0.2 kb selectable tRNA suppressor gene (sup Phe), and a 0.4 kb cer (ColE1 resolution) fragment to resolve pCOR oligomers.

Such plasmids can only replicate in π-producing bacteria considerably limiting their host range. pCOR selection uses expression of a synthetic amber suppressor tRNA gene, specific for phenylalanine (sup Phe), which does not require antibiotics. This suppressor corrects an amber mutation in argE gene making it possible for the recombinant host strain to grow on a minimal medium lacking arginine (Figure 1).

pCOR host vector system. The XAC-1pir116 strain requires arginine for growth on minimal medium whereas the arginine deficiency of the pCOR host is corrected by the tRNA sup Phe from the plasmid. The pCOR plasmid, the pir116 and argEam genes, and the XAC-1pir116 chromosome are not drawn to scale. The amber mutation is shown by a stop and its correction by a crossed-out stop. XAC-1pir116 genotype: ara Δ(lac-proB)x111 gyrA argEam rpoB thi uidA(ΔMluI)::pir116 F′(proB+ lacI373 lacZu118am).

The pCOR conditional origin of replication comes from a natural plasmid, R6K, rather than the widely used ColE1-derived plasmids. R6K is a 38 kb theta-replicating R (resistance) plasmid. R6K replication functions5 are clustered in a 5.5 kb DNA fragment that contains three origins of replication: α and β that are mainly used, and γ. All three are under the control of a single replication initiation protein, π, the product of the pir gene. This protein binds to direct nucleotide sequence repeats (seven tandemly associated 22-bp direct repeats) at the origin of replication. It is a dual regulator, exerting both a positive effect on replication and negative feedback that maintains R6K at its characteristic copy number (15 per genome). Copy-up mutants inhibit replication less and maintain the γ origin at a high copy number.pir116 6 contains a single base mutation (CCC→CTC) that leads to a proline to leucine substitution at position 106 in π.

The minimum genetic information required to maintain R6K derivatives is the 400-bp γ origin (ori γ) and the cis or trans-acting π initiator protein.5 Ori γ was isolated from the suicide vector pUT-T7pol7 as a 0.4 kb BamHI–EcoRI fragment and was ligated to the kanamycin resistance gene (Km R ) from pUC4K8 to generate pXL2666. Ori γ replication and plasmid copy number were assayed in wild-type and copy-up backgrounds. pXL2666 was transferred into a pir (BW190949) and a pir116 strain (BW196109) of E. coli. Agarose gel electrophoresis showed that there were five to 10 more plasmid DNA copies in the pir116 than in the pir strain, consistent with previous work6 (data not shown). It also pointed out that the monomer/oligomer ratio of R6K-derivatives was affected by the presence of the copy-up mutation which resulted in the major plasmid species being multimers, mostly dimers (Figure 2). Plasmid linearization by EcoRI indicated that these preparations contained a mixture of monomer and the corresponding oligomers. Topoisomerase treatment showed that the high molecular weight species were circular, supercoiled, plasmid multimers rather than catenanes. These plasmids are relaxed by DNA topoisomerase I and topoisomerase IV10 and no decatenation by topoisomerase IV was observed, although this enzyme was active on kinetoplast DNA (data not shown). Plasmid multimerization is not compatible with pharmaceutical use due to low lot to lot reproducibility and too few molecules per unit weight. As well as reducing the number of independently segregating molecules, it may also severely reduce plasmid stability by affecting plasmid segregation during bacterial cell division. In our effort to resolve multimers, we tested the cer fragment that can promote monomerization independently of the replicon involved. This cis-acting DNA determinant reduces multimerization via a site-specific recombination event that requires only E. coli-encoded proteins. Two recombinases, XerC and XerD, are necessary, along with two accessory proteins, ArgR and PepA, that ensure exclusively intramolecular recombination at the cer site.11 The 0.38 kb HpaII cer fragment12 from the E. coli plasmid ColE1 was inserted into pXL2666. Comparison of pXL2666 (ori γ-Km R ) and pXL2730 (ori γ-Km R -cer) topology (Figure 2) showed that cer leads to monomerization of R6K-derived plasmids. A deletion12 leading to the inactivation of cer, was introduced into pXL2666 to generate pXL2754 (ori γ-Km R -Δcer). This construct led to the production of plasmid multimers. These experiments were also carried out in XAC-1pir and XAC-1pir116 strains (data not shown) which differ only by the pir allele. Thus, these results shed light on the link between plasmid multimerization/copy-up pir mutations and multimer resolution via cer-mediated intramolecular recombination.

Influence of cer on the topology of R6K-derived plasmids. (a) Plasmid constructs. B: BamHI E: EcoRI M: MluI restriction sites. (b) Agarose gel analysis of plasmid topology. Plasmids were prepared by CsCl-ethidium bromide gradient and analyzed by agarose gel electrophoresis. 1, pXL2666 2, pXL2754 3, pXL2730. a, linear (EcoRI digested) plasmid b, circular plasmid.

An important original feature of pCOR is the absence of an antibiotic resistance gene for selection in E. coli. E. coli amber (TAG) suppressor tRNA genes do not contain any sequences derived from phages or transposons and are very small (less than 100 bp). They are efficient plasmid selection markers, even in high-density cultures of E. coli.13 A synthetic amber suppressor tRNA gene, specific for phenylalanine (sup Phe) was used as the pCOR marker. sup Phe is expressed from a strong constitutive E. coli promoter, Plpp. This cassette (0.17 kb NarI–HindIII fragment) was isolated from pCT2-F14 and was tested in E. coli XAC-1.14 The chromosomal argE gene of XAC-1 contains an amber mutation that results in a truncated and therefore inactive product. As argE encodes N-acetylornithinase which is essential for arginine biosynthesis, this strain cannot grow in the absence of arginine. The suppressor tRNA sup Phe causes the correct translation of mRNA and the synthesis of an active full-length N-acetylornithinase. Thus, XAC-1 can grow on minimal medium lacking arginine as long as the plasmid is present, ensuring plasmid selection. However, this strain does not synthesize the R6K π initiator protein and thus cannot replicate pCOR. Therefore, the π-encoding gene, pir (or the copy-up mutant pir116), was inserted into the XAC-1 chromosome by homologous recombination. This method does not introduce any mobile genetic element, such as transposons or phages, that affect genome stability. Since there is no similarity between pir and the bacterial genome, pir or pir116 were inserted into the chromosomal uidA gene which encodes β- D -glucuronidase (EC, as described by Metcalf et al.9 This gene provides sufficient sequence similarity with the E. coli genome for recombination to occur. β-Glucuronides are not essential in standard growth conditions, therefore it is possible to inactivate uidA by replacement with the gene of interest (pir or pir116). Replacement of the genomic uidA gene with the uidA::pir116 cassette which results in uidA inactivation, is easy to detect with a chromogenic substrate, X-gluc, as the β-glucuronidase-positive colonies develop a blue color. The Km R -uidA:: pir116 cassette9 from M13wm33 (or pir from M13wm34) was inserted into the BamHI site of the M13mp1015 replicative form. This recombinant bacteriophage is a suicide vector in non-suppressor E. coli strains. The gene replacement technique developed by Blum et al16 was successful in 18–34% of the deoxycholate-resistant and kanamycin-sensitive segregants as demonstrated by the loss of β-glucuronidase activity. These clones were tested by assessing replication of an R6K derivative, pBW30.9 The integrity of the bacterial genome in the region close to the site of homologous recombination was analyzed by PCR. Each unique amplified fragment, all of which were of the expected size, was identified by restriction analysis. The other genetic markers argEam, lacZam, gyrA (nalidixic acid resistance), rpoB (rifampicin resistance), presence of episomal F′ (susceptibility to M13 infection), were unaffected. The resulting pCOR host strains were named XAC-1pir and XAC-1pir116. A pCOR plasmid (pXL2760: ori γ-Km R -cer-sup Phe) was introduced into these strains and subsequent selection by plating on minimal medium demonstrated that these two strains allow pCOR replication and selection. The yield of pCOR from a small-scale culture of the XAC-1pir116 recombinant strain was similar to that of standard plasmids such as pUC and restriction analysis pattern was found to conform to theory (data not shown).

Since plasmid DNA degradation could occur during the production/purification process, a pCOR host lacking endonuclease I was constructed. Endonuclease I, encoded by endA, is a periplasmic DNase which is inhibited by RNA and internally cleaves native DNA into oligodeoxyribonucleotides in a sequence-independent fashion.17 This enzyme is not essential and its biological role in E. coli is still unknown as mutants resemble wild-type cells in terms of growth rate, phage propagation and conjugation properties.18 Various studies19,20 have found that EndA − strains produce DNA of higher quality than EndA + E. coli strains. A DNA fragment containing endA21 was used to construct an M13 Km R suicide vector. The 0.37 kb FspI–PvuII central fragment was deleted removing 52% of endA. Gene replacement of the wild-type endA by the deleted endA was then obtained as previously described for the construction of XAC-1pir116. Genomic modification of the locus was verified by PCR and the absence of endonuclease I activity was demonstrated by two biochemical tests. First, gene replacement was monitored by a methyl green staining assay.17 Second, the absence of endonuclease activity was assayed using the breakage of supercoiled DNA in the presence of cold-shock fluid22 from mutant or wild-type strains. Plasmid DNA was totally degraded with an extract from XAC-1pir116 (EndA + strain) whereas no degradation was observed with the EndA − isogenic strain (XAC-1pir116 endA − ).

XAC-1pir116 contains an F′ episome, a circular DNA molecule of approximately 100 kb, that carries proB + lacI373 lacZu118am. Many male E. coli laboratory strains carry a traD36 mutation on their episome, but no mutation affecting F′ transfer ability has been described for XAC-1. traD is at the 5′ end of one of the tra (transfer) operons and encodes a membrane protein that is directly involved in DNA transfer and DNA metabolism23 furthermore, traD mutants do not transfer DNA.24 The presence of wild-type traD can be considered as a potential risk of plasmid dissemination between E. coli strains by nonspecific transfer of co-resident plasmids.25 Therefore, the episomal traD gene of XAC-1pir116 was inactivated to abolish F′ transfer. A 2 kb central fragment from traD, comprising 92% of the gene, was replaced with the 2 kb omega element from pHP45Ω26 by homologous recombination in XAC-1pir116 endA − . The omega element contains the aadA antibiotic resistance gene flanked by short inverted repeats. aadA encodes aminoglycoside-3 adenyltransferase and confers resistance to streptomycin and spectinomycin. The omega fragment was used because it prematurely terminates RNA and protein synthesis leading to the inactivation of the whole traD operon, not as a selectable marker (no antibiotic added during plasmid production). PCR and restriction analysis of the modified traD locus showed the expected pattern of amplified fragments demonstrating that gene replacement had occurred. The resultant strain was named TEX1. This new pCOR host strain, which is more suitable for industrial applications, is being evaluated.

Since gene therapy applications require large amounts of plasmid DNA, we assessed the pCOR host strain in fermentation experiments. Complex media containing yeast extract were used for fed-batch fermentation with XAC-1pir116. pCOR stability (more than 50 generations) makes it possible to use such non-selective media. In these conditions, XAC-1pir116 produced more than 40 g/l dry cell weight and 100 mg/l of pCOR pXL2774 (described in Figure 3) could be obtained from 2-liter fermentors (Table 1). pCOR copy number was evaluated at 400–500 plasmids per cell and the rate of plasmid DNA synthesis was constant throughout fermentation. These results were extrapolated to an 800-liter fermentor suitable for manufacturing (Table 1). To control culture conditions further, fermentation was also performed in the absence of yeast extract or any raw material from animal origin. Similar results (30 g/l dry cell weight and 100 mg/l of plasmid DNA) were obtained using a defined medium in 2-liter cultures with no loss of productivity (Table 1).

In vivo transfection with pCOR-Comparison of luciferase vectors after injection into mouse muscle. Tibialis anterior muscles of mice were injected percutaneously with 30 μl of DNA in saline containing 3 × 10 12 luciferase expression cassettes (approximately 15 μg of naked DNA). Mice were killed 72 h after injection. The muscles were collected and transferred to the lysis solution from the Luciferase assay system (Promega, Madison, WI, USA). The extraction was further performed according to the manufacturer’s instructions. The luciferase activity was measured using a LUMAT LB9501 luminometer (Berthold, Evry, France). Results are indicated as nanograms (ng) of luciferase per injected muscle as mean ± standard error of the mean. Plasmid DNA was purified by CsCl-ethidium bromide gradient ultracentrifugation and analyzed by agarose gel electrophoresis. Endotoxin level ranged between 1 and 5 EU/mg of plasmid DNA. The major structural elements of the plasmids vectors are indicated. The name of the pCOR plasmids is underlined. CMV, human cytomegalovirus immediate–early1 enhancer/promoter SV40, simian virus 40 promoter TK leader, leader sequence from HSV1 TK gene luc, peroxisomal luciferase gene luc+, modified (cytoplasmic) luciferase gene SV40 late/early, simian virus 40 (SV40) late/early polyadenylation signal bGH, bovine growth hormone polyadenylation signal.

pCOR gene transfer in vitro and in vivo was assessed and compared with commercial plasmids. Various pCOR plasmids were constructed with luc, the gene encoding firefly luciferase, or luc+, the engineered cytoplasmic luciferase gene. These plasmids were efficient in standard in vitro transfection of murine or human cell lines, giving higher levels of reporter gene activity than commercial plasmids (ie pGL3-basic Promega, Madison, WI, USA) (data not shown). The same amount of luciferase expression cassette (3 × 10 12 ) in plasmids pGL3-control, pXL2774 and pXL3133 gave 0.07, 0.4 and 16.5 ng of luciferase per muscle (n = 24), respectively, when injected into the tibialis anterior of OF1 mice, as determined 72 h after injection (Figure 3). Thus the pCOR plasmid pXL3133 with an improved luciferase expression cassette is efficient at driving expression in muscle with levels of reporter gene activity at least 200-fold higher than that of a reference commercial vector. This result was comparable with those previously described with optimized ColE1-derived luciferase reporter plasmids.27

Nonviral gene therapy has been based exclusively on E. coli plasmids derived from the ancestral ColE1 plasmid. These plasmids replicate in the absence of plasmid-encoded proteins and their maintenance only requires E. coli host-encoded factors, such as DNA polymerases I and III and DNA-dependent RNA polymerase. To restrict plasmid host range to a single E. coli strain, we designed a plasmid derived from an E. coli natural narrow-host range plasmid,28 R6K. Unlike ColE1 replication, R6K requires synthesis of the plasmid-coded protein π that controls R6K replication through ori γ core sequences. pCOR replication is restricted to a specific bacterial host expressing π. The γ origin from R6K has no identified open-reading frame, no undesirable sequences such as insertion sequences, and no element likely to direct its transfer to other bacteria. Although R6K contains two transfer origins,29 they are not present in the γ origin of replication. Bacterial host F′ episome transfer was abolished by insertional inactivation of the episomal traD to avoid low efficiency illegitimate co-transfer of pCOR.

The pCOR bacterial hosts, XAC-1pir116 or derivatives such as TEX1, are specifically engineered E. coli strains containing a chromosomal copy of a copy-up variant of pir, pir116, inserted into the nonessential uidA gene. R6K derivatives have a low copy number (around 15 copies per genome) which is not compatible with industrial-scale production. Introduction of pir116 into the XAC-1 genome generates high copy number pCOR where plasmid multimerization was resolved by inserting the cer resolution site into pCOR. This is the first report of plasmid multimers in pir116 strains and of cer-mediated multimer resolution. We successfully cloned inserts of 8 kb into the pCOR backbone without any effect on plasmid copy number or stability indicating that pCOR or ColE1 derivatives appeared to have similar cloning capacity. R6K derivatives can accommodate inserts as large as 50–100 kb in a copy up context.30

R6K γ origin-bearing plasmids are widely used as suicide vectors in various bacteria, such as E. coli (non-pathogenic9 or pathogenic31 strains), Yersinia pestis,32 Pasteurella,33 Rhodobacter spheroides,34 Pseudomonas putida or Klebsiella pneumoniae.7 This strongly supports the fact that pCOR replication is totally dependent on π initiator protein, and is thus biologically safe. The replication of natural R6K, which encodes π, is restricted to E. coli and closely related bacteria.28 To our knowledge, there is no report of a π-like activity that may allow the replication of R6K derivatives. The potential risk of dissemination of pCOR would be restricted to a bacterium containing a resident R6K. However, incompatibility35 between plasmids that share the major incompatibility factor36 present in ori γ will lead to the loss of one of these plasmids and in any case, will result in only ephemeral pCOR replication since there is no selective advantage for bacteria to maintain a pCOR. pCOR replication is conditional and offers environmental advantages over current plasmids used in gene therapy.

Plasmids are maintained as stable extrachromosomal DNA molecules and are excellent vectors for the dissemination of antibiotic resistance genes37 to a wide range of bacteria. Contamination of the final product intended for use in gene therapy with an antibiotic could cause allergic reactions. The use of a suppressor tRNA as a selectable marker in bacteria is therefore an interesting alternative to the commonly used drug resistance markers. This tRNA, for which the anticodon was mutated to match the amber termination triplet, mediates plasmid selection in bacteria by suppression of an amber mutation in a gene essential for growth in the conditions used. This system developed by Normanly et al14 proved to be efficient even in fermentation experiments.13 An amber suppressor tRNA may cause readthrough at a significant number of natural termination codons and induce abnormal protein synthesis. However, this requires the conjunction of several events in mammals: (1) sufficient transcription of the suppressor tRNA in mammalian cells (2) correct folding of this tRNA which is essential to its activity (3) correct recognition and (4) loading of a specific prokaryotic tRNA by a eukaryotic tRNA synthetase. Several experiments suggest that this is unlikely. Jacobson38 obtained no aminoacylation of E. coli wild-type tRNA Phe with extracts from rat liver, mouse liver, maize or human spleen. It has also been shown that even if an E. coli suppressor tRNA Gln is expressed in mammalian cells, amber suppression is dependent upon co-expression of the E. coli GlnRS aminoacyl-tRNA synthetase gene.39 Expression of sup Phe can only result from unknown cryptic eukaryotic expression signals in the plasmid backbone. To avoid spurious transcription, the eukaryotic expression cassette was inserted into pCOR such that the transgene and sup Phe gene are in opposite orientations.

Amber suppression is never complete and most known mammalian genes (79%40) terminate in UAA or UGA. Consistent with these observations, natural mammalian amber suppressor tRNA were isolated from calf liver.41 We detected no apparent changes in cell viability when pCOR were transfected in murine or human cell lines in vitro or in murine muscle cells in vivo. Furthermore, when eukaryotic amber suppressor tRNA are expressed from appropriate eukaryotic expression signals, no adverse effects are noticed in transfected mammalian cells42,43 or in whole plants.44 Thus, the bacterial suppressor tRNA Phe is unlikely to cause any deleterious effects in human cells and is an efficient selectable marker for pCOR in E. coli.

Short (6 bp) immunostimulatory sequences (ISS) in bacterial plasmid DNA have been shown to trigger powerful immune responses and to affect gene expression in mice.45 Schwartz et al46 found that unmethylated CpG motifs caused inflammation of the lower respiratory tract of mice that received DNA by intratracheal instillation. Results obtained with oligonucleotides47 gave indication of ISS potential to stimulate human B cells but the effects in man of plasmidic ISS have still to be investigated. However, the results obtained in mice suggest that these sequences may also cause problems in humans. The pCOR backbone does not contain ISS. Moreover, there are no ‘two 5′ purines – unmethylated CpG – two 3′ pyrimidine’ motifs in the pCOR backbone. In contrast, a plasmid containing the widely used Tn903 kanamycin resistance gene from pUC4K (Pharmacia Biotech, Uppsala, Sweden), for example VR1255,27 and the ColE1-derived origin of replication from pBlueScript (Stratagene, La Jolla, CA, USA) contains nine such motifs. Thus pCOR should have less, if any, immunostimulatory properties than standard ColE1-derived plasmids. If DNA vaccination is envisaged, immunostimulatory sequences could be added to the plasmid backbone.

The pCOR backbone is significantly smaller (1 kb) than standard ColE1-derived plasmids (2.5 kb). It contains a minimal number of prokaryotic DNA sequences, the functions of which are known. The reduction of the amount of prokaryotic sequences in plasmid vectors is of importance since some of these sequences were shown in different studies27,28,29,30,31,32,33,34,35,36,37,38,39,40,41,42,43,44,45,46,47,48 to interfere with eukaryotic gene expression.

We aimed to improve biosafety during plasmid production and nonviral gene therapy by totally redesigning the plasmid backbone. The pCOR plasmid and host system are suitable for clinical and industrial applications in terms of safety (conditional origin of replication, no antibiotic marker, minimal plasmid sequences, inactivation of bacterial episome transfer) and gene expression (at least as efficient in vitro and in vivo as standard ColE1-derived plasmids, smaller size). In addition, pCOR has proved to be appropriate for production (high yield: 100–150 mg/l in fermentors, possibility of using defined media) and purification (small size, high copy number, elimination of endonuclease I activity of the host).

In conclusion, pCOR is an appropriate DNA vector for gene delivery which offers environmental advantages over current plasmids without any loss in gene transfer efficiency.