Molecular Biotechnology: Principles and Applications - Glick B., Pasternak J. 2002

Fundamentals of Molecular Biotechnology
Recombinant DNA Technology
Plasmid Vectors

Plasmids are extrachromosomal, autonomously replicating, double-stranded circular DNA molecules. They are found in virtually all Bacteria. Some plasmids carry information that ensures their own transfer from one Cell to another (F plasmids), while others carry Antibiotic Resistance genes (R plasmids) or specific sets of genes responsible for the utilization of unusual metabolites (degradation plasmids). Additionally, there are plasmids in which no genes with clearly defined Functions have been detected (cryptic plasmids, derived from the English term 'cryptic', meaning hidden or latent). Plasmid sizes range from less than 1 to over 500 kbp. Each plasmid contains an origin of Replication (ori) site, without which replication within the host cell would be impossible.

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Fig. 4.6. T4 DNA ligase forms phosphodiester bonds between 5'-phosphate and 3'-hydroxyl groups at a nick in the double-stranded DNA backbone. A. Ligation of sticky ends. B. Ligation of blunt ends.

Some plasmids are present in The Cell in 10–100 copies; these are referred to as high-copy-number plasmids. Low-copy-number plasmids are present at 1–4 copies per cell. Plasmid DNA typically accounts for 0.1–5.0% of the total cellular DNA. If two or more plasmids cannot coexist within the same cell, they are said to belong to the same incompatibility group. Plasmids belonging to different incompatibility groups coexist freely in the same cell, regardless of their copy number. In some microorganisms, up to 8–10 different plasmids have been found in a single cell, with each performing its own functions, maintaining its characteristic copy number, and belonging to its own incompatibility group. Some plasmids carry a specific replication initiation site and can replicate only in Cells of a single species. In other plasmids, this site is less specific, allowing them to replicate in A wide variety of bacterial cells. Accordingly, plasmids are classified as having either a narrow or a broad host range.

As autonomously replicating genetic elements, plasmids possess all the essential properties that make them suitable as vectors for transferring cloned DNA. However, natural (unmodified, non-engineered) plasmids quite frequently lack certain features mandatory for a 'high-quality' vector. These important properties include:

1) a small size, since the efficiency of exogenous DNA transfer into E. coli decreases significantly when the plasmid length exceeds 15 kbp;

2) the presence of a unique restriction site where insertion can be carried out; 3) the presence of one or more selectable genetic markers to identify recipient cells carrying recombinant DNA. Therefore, plasmid vectors must be engineered using Recombinant DNA technology.

The pBR322 Plasmid Vector

In the 1980s, the pBR322 plasmid vector was one of the most popular universal vectors. Typically, the designation of a plasmid vector includes the lowercase letter p (for plasmid) and several additional letters related to the vector's description or history of creation. For instance, the letters BR in the designation of plasmid pBR322 indicate the authorship of F. Bolívar and R. Rodríguez, who constructed this plasmid, and the number 322 is a numerical designation taken from their research protocols. The length of plasmid pBR322 is 4361 bp. It carries two antibiotic resistance genes (Fig. 4.7), ampicillin (Amp^r) and tetracycline (Tet^r), as well as unique sites for BamHI, HindIII, and SalI within the Tet^r Gene, a single PstI site within the Amp^r gene, a single EcoRI site located outside the coding sequences, and an origin of replication that ensures replication exclusively in E. coli. The plasmid replicates to a high copy number and is transferred to other bacterial cells with difficulty. How does the pBR322 cloning vector work? If purified circular pBR322 plasmid is treated with a restriction endonuclease that cuts it at a single site located within one of the antibiotic resistance genes, a linear molecule with sticky ends is produced. These molecules are mixed with donor DNA containing the desired gene, which has been pre-treated with the same restriction enzyme. Because the sticky ends of these two DNAs are mutually complementary, they anneal to form hybrid molecules. Next, the mixture is treated with T4 phage DNA ligase in the presence of ATP, resulting in a multitude of different fragment combinations as well as undesired products, notably self-joined donor DNA fragments and original plasmid DNAs. To reduce The amount of the latter, the restricted plasmid DNA is treated with alkaline phosphatase, which removes 5'-phosphate groups from the linearized molecule: DNA ligase cannot seal the ends of dephosphorylated linear plasmid DNA (Fig. 4.8). As for the recombinant DNA molecules themselves, although they contain two single-stranded nicks, their fragments are held together by two phosphodiester bonds formed via DNA ligase between the dephosphorylated plasmid DNA and the restricted donor DNA (Fig. 4.8). Following replication in the transformed cell, the single-stranded nicks are repaired by the host cell's ligation system.

Fig. 4.7. Genetic Map of the pBR322 plasmid vector. The tetracycline (Tet^r) and ampicillin (Amp^r) resistance genes contain unique recognition sites for HindIII, SalI, BamHI, and PstI. The EcoRI site is located outside these genes. The vector length is 4361 bp.

Transformation and Selection

Next, the recombinant DNA must be introduced into the host cell. This process is called transformation. To achieve this, specially developed techniques are used, such as exposing cells to heat Shock and treating them with calcium chloride (CaCl₂). However, transformation efficiency remains relatively low; typically, no more than one in a thousand cells is transformed. Thus, the majority of cells after transformation do not contain recombinant DNA. Some of them acquire rejoined circular plasmid DNA that escaped dephosphorylation by alkaline phosphatase, others take up non-plasmid DNA, and only a few contain a plasmid with an integrated foreign DNA insert (hybrid plasmid).

Fig. 4.8. Insertion of foreign DNA into a plasmid vector. Plasmid DNA treated with a restriction enzyme and alkaline phosphatase is mixed with restricted donor DNA containing the target gene, and DNA ligase is added. Two of the four single-stranded nicks are thereby sealed, and the construct is stabilized by the newly formed phosphodiester bonds. After introducing the hybrid DNA into the host cell, it replicates, forming new circular molecules that are nick-free.

As mentioned earlier, extrachromosomal DNA lacking an origin of replication cannot replicate in a bacterial cell. Thus, The entry of exogenous DNA into a cell does not guarantee that it will be maintained within the host cell. Furthermore, to preserve the recombinant DNA in its original form within the host cell, it is necessary that the cell lacks genes encoding restriction Enzymes that could lead to its degradation, and that the cell has a RecA^- phenotype (such cells are incapable of general recombination, preventing exogenous DNA from being modified through Homologous Recombination).

Subsequently, it is necessary to identify cells containing recombinant DNA. The identification method must be as simple as possible because a vast number of Cells must be screened. In the pBR322 system, where foreign DNA is inserted into the BamHI site, specific identification involves two stages. First, the transformed cells are plated onto a nutrient medium containing ampicillin. Under these conditions, only cells harboring an intact Amp^r gene can grow—either as part of an intact pBR322 plasmid or as part of a hybrid plasmid; untransformed cells are sensitive to ampicillin. The BamHI site is located within the Tet^r gene of plasmid pBR322 (Fig. 4.7); insertion of a DNA fragment into this gene interrupts the coding sequence, leading to the loss of tetracycline resistance. Consequently, cells carrying the hybrid plasmid are resistant to ampicillin but sensitive to tetracycline, whereas cells that received the intact pBR322 plasmid carry the Tet^r gene and are resistant to both ampicillin and tetracycline.

In the second stage, these two variants are separated. Cells that grew on the ampicillin medium are replica-plated onto tetracycline medium. Cells forming colonies on the tetracycline plates contain the intact pBR322 plasmid because, as noted earlier, they are resistant to both ampicillin and tetracycline. Cells that fail to grow on the tetracycline plates are sensitive to this antibiotic, meaning they harbor the recombinant pBR322 plasmid.

Among the colonies that grew on the ampicillin medium, those found to be sensitive to tetracycline are isolated, and individual cell clones are derived from each colony, or—more commonly—all ampicillin-resistant, tetracycline-sensitive colonies are pooled and cultured together. Further screening can then be performed to identify cells carrying the pBR322 hybrid plasmid with the specific insert. The presence of HindIII and SalI sites within the Tet^r gene and a PstI site within the Amp^r gene of plasmid pBR322 allows for alternative cloning strategies for foreign DNA fragments. If the PstI site is used for insertion, selection follows the same logic but in reverse order: cells are first plated on tetracycline medium and then on ampicillin medium.

Other Plasmid Vectors

The idea of using pBR322 as a cloning vector was quite successful, but this plasmid contains only a limited number of restriction sites, and screening transformed cells is time-consuming. This led to the need to develop alternative cloning systems. For example, the 2686 bp plasmid pUC19 contains: an ampicillin resistance gene; a regulatable segment of the E. coli lactose Operon ß-galactosidase gene (lacZ'); the lacI gene encoding the repressor that controls lacZ' Gene Expression; a polylinker—a short sequence containing multiple unique recognition sites for endonucleases (EcoRI, SacI, KpnI, XmaI, SmaI, BamHI, XbaI, SalI, HincII, AccI, BspMI, PstI, SphI, and HindIII); and THE ORIGIN OF replication from plasmid pBR322 (Fig. 4.9).

Fig. 4.9. Genetic map of the pUC19 plasmid vector. The plasmid consists of 2,686 Base Pairs and contains unique recognition sites for EcoRI, SacI, KpnI, XmaI, SmaI, BamHI, XbaI, SalI, HincII, AccI, PstI, BspMI, SphI, and HindIII localized within the polylinker; an ampicillin resistance gene; a replication initiation site functioning in E. coli; and the lacI gene, which controls the synthesis of a repressor that blocks lacZ' gene METABOLISM/31.html">Transcription in the absence of the IPTG inducer.

Selection of transformed cells is based on the following principles. If cells containing the unmodified pUC19 plasmid are grown in the presence of isopropyl-ß-D-thiogalactopyranoside (IPTG), an inducer of the lac operon, the lacI gene product cannot bind to the promoter-operator region of the lacZ' gene, resulting in the transcription and Introduction/27.html">Translation of the plasmid lacZ' fragment. This fragment product interacts with the protein encoded by the chromosomal DNA, yielding active ß-galactosidase. The multiple cloning site (polylinker) is inserted into the lacZ' gene without disrupting The production of functional ß-galactosidase; therefore, if its substrate 5-bromo-4-chloro-3-indolyl-ß-D-galactopyranoside (X-Gal) is present in the medium, it is hydrolyzed by this enzyme to produce a blue-colored product that stains colonies of cells containing the unmodified pUC19 plasmid.

To clone DNA into pUC19, donor DNA is cleaved with a restriction enzyme whose recognition site is located within the polylinker; plasmid DNA is digested with the same restriction enzyme and then treated with alkaline phosphatase. Both DNA preparations are mixed in the presence of T4 DNA ligase, and the resulting product is used to transform cells capable of synthesizing the fragment of ß-galactosidase (LacZa) that combines with the lacZ' gene product to form an active enzyme. The treated cells are plated onto a nutrient medium supplemented with ampicillin, IPTG, and a ß-galactosidase substrate. Nontransformants cannot grow in the presence of ampicillin, whereas cells carrying the intact plasmid form blue colonies on ampicillin-containing media. Host cells harboring a recombinant plasmid form white colonies on the same medium, because insertion of foreign DNA into the polylinker typically disrupts this sequence. This issue can be resolved by using a different restriction enzyme.

The step following library construction is the screening process to identify the clone or clones carrying the target DNA sequence. Three widely used Methods are employed for this purpose: Hybridization with a labeled DNA probe followed by autoradiographic analysis, immunological screening, and screening for The activity of the protein encoded by the target gene.

Hybridization Screening

A desired nucleotide sequence in a DNA sample can be detected using a DNA probe that specifically pairs only with the target sequence. To achieve this, the DNA is first converted into single-stranded form by heat Treatment or exposure to alkali. Under these conditions, the Hydrogen Bonds between the bases break and the strands separate (Denaturation). If the Temperature is then slowly lowered, the strands will reassociate (renaturation). If a single-stranded DNA probe is present in the solution during this process, it will also renature with the DNA by specifically pairing with complementary regions. This results in The formation of a hybrid DNA molecule, i.e., a double-stranded molecule whose strands originate from two different DNA sources.

Fig. 4.11. DNA hybridization. The target DNA is denatured and immobilized on a solid support, such as a nitrocellulose or nylon filter. A labeled DNA probe (typically 100 to 1,000 bp in length) is also denatured, applied to the filter containing the target DNA, and allowed to anneal. To remove unbound DNA probe, the filter is washed and the label is visualized. If no hybridization occurs between the probe and the target DNA, no signal is detected on the filter. (The label is indicated by a colored asterisk in the figure.)

The DNA hybridization Procedure is carried out as follows. The target DNA is denatured, and the single-stranded molecules are irreversibly bound ("fixed") to a solid support (a nitrocellulose or nylon filter). This immobilization step is typically performed at high temperature. The filter is then incubated with a single-stranded DNA probe labeled with a radioisotope or another reporter group. If The nucleotide sequences of the probe and the target DNA are complementary, they pair (i.e., hybridize) (Fig. 4.11). The hybrid molecules can be visualized by autoradiography (Box 4.2) or another method depending on The Nature of the label. If there is no complementarity between the probe and the target DNA, hybridization does not occur, yielding a negative result. Typically, the probe size ranges from 100 to 1,000 bp or more, although both larger and smaller probes can be used. For successful hybridization—that is, the formation of a stable complex—it is generally required that over 80% of the NUCLEOTIDES match across a 50-nucleotide stretch, although this depends on the stringency of the reaction conditions.

Labeled DNA probes can be generated in several ways. One approach, known as the random priming method, utilizes a mixture of synthetic oligonucleotides (oligomers) containing all possible hexanucleotide combinations. Some of these oligonucleotides are complementary to sequences within the target DNA and hybridize to them after the DNA has been denatured (Fig. 4.12). Following the annealing of the oligonucleotides to the denatured DNA template, four deoxynucleotides (deoxynucleoside triphosphates; dNTPs)—one of which is labeled—and the Klenow fragment of E. coli DNA polymerase I are added to the reaction mixture. The Klenow fragment possesses DNA polymerase and 3'-exonuclease activities, but lacks the 5'-exonuclease activity characteristic of intact E. coli DNA polymerase I, which would otherwise degrade newly synthesized DNA molecules. The single-stranded Regions of the target DNA serve as templates for the synthesis of new DNA molecules, while the randomly bound oligonucleotides act as primers (Fig. 4.12). In radioactive labeling, one of the dNTPs contains a^-32P, rendering the probe itself ³²P-labeled. The radioactive signal is subsequently detected via autoradiography.

Fig. 4.12. Synthesis of a labeled DNA probe by random priming. Hexanucleotides (a mixture of all possible hexamer combinations) are added to denatured double-stranded DNA containing The nucleotide sequence to be used as a probe, and the mixture is annealed. Some of the oligonucleotides hybridize to the unlabeled denatured DNA and, in the presence of the Klenow fragment and four dNTPs (one of which is labeled [*]), serve as primers for complementary strand synthesis. Following denaturation of the newly synthesized DNA, a mixture of labeled DNA fragments is obtained, which collectively span virtually the entire original DNA template.

A common non-radioactive label is biotin, which is chemically linked to one of the four dNTPs. To detect the hybridized biotinylated probe, the filter is treated with a streptavidin-enzyme conjugate (such as alkaline phosphatase). Streptavidin forms a high-affinity complex with biotin, which is then visualized by supplying a substrate that the enzyme converts into a colored or luminescent product.

Probes for screening Genomic Libraries can be obtained by at least two distinct approaches. First, cloned DNA from a closely related Organism can be used (a heterologous probe). In this case, hybridization conditions must be optimized to allow pairing despite substantial sequence divergence between the probe and the target DNA, thereby overcoming discrepancies between the source DNA of the probe and the DNA under investigation. Second, a probe can be chemically synthesized based on the known Amino Acid Sequence of the protein product encoded by the target gene.

Genomic DNA libraries are typically screened According to the following protocol. Following transformation, cells are plated on nutrient Agar, and the resulting colonies are transferred to a solid support (such as a nitrocellulose or nylon filter); the cells are then lysed, and the released DNA is deproteinized, denatured, and fixed to the support. A labeled probe is applied to the filter to allow annealing, followed by autoradiography. Colonies on the original master plate corresponding to those containing the hybridized DNA are isolated and cultured (Fig. 4.13). Because most libraries are generated via partial Digestion, a positive hybridization signal may be obtained for multiple colonies (clones). It is then necessary to determine which specific clone, if any, contains the complete gene. Gel Electrophoresis and restriction mapping are used to determine the size of each insert and to identify identical or overlapping fragments. Alternatively, additional cloning can be performed to assemble the full-length gene from overlapping fragments. If the insert in a particular clone is sufficiently large to encompass the entire gene, it can be sequenced to verify the presence of start and stop codons as well as the complete Open Reading Frame encoding the target protein.

Unfortunately, there is no guarantee that a given library contains the entire nucleotide sequence of the gene of interest. If the search for a full-length gene is unsuccessful, a new library can be constructed using a different restriction enzyme and screened using the original probe or probes derived from the previous library. To increase the likelihood of capturing the complete gene, one can also construct libraries containing DNA fragments substantially larger than the average prokaryotic gene (an approach we discuss later in this chapter).

Immunological Screening

In the absence of a DNA probe, alternative methods can be used to screen a genomic library. For instance, if the cloned gene is expressed, its protein product—either in full or in part—can be detected using immunological techniques. Technically, this procedure shares many similarities with nucleic acid hybridization. All clones of the library are plated on nutrient medium, and the resulting colonies are transferred to a filter, where the cells are lysed and the released Proteins are immobilized. Primary Antibodies that specifically bind to the protein of interest (antigen) are then applied to the filter. After washing away unbound primary antibodies, the filter is incubated with secondary antibodies directed against the primary antibodies. Many detection systems employ enzyme-conjugated secondary antibodies, such as those linked to alkaline phosphatase. Following a wash step, a colorless substrate is added. If the secondary antibodies have bound to the primary antibodies, the enzyme catalyzes the Hydrolysis of the substrate, generating a colored precipitate at the site of the reaction (Fig. 4.14).

Fig. 4.13. Screening a genomic library using a labeled probe. Transformed cells are plated onto a solid nutrient medium that selects for transformants. 1. Cells from each grown colony are transferred to a solid support (e.g., a nitrocellulose or nylon filter) in a pattern that replicates their positions on the plate. 2. The cells are lysed, and the released DNA is denatured, deproteinized, and fixed to the filter. 3. A labeled DNA probe is applied to the filter to allow hybridization. Unbound probe is washed away, and autoradiography is performed to identify which cells have bound the labeled probe. 4. Colonies on the master plate containing the target DNA (yielding a positive hybridization signal) are identified, picked, and cultured.

The cells on the plate that correspond to the colored spots on the filter contain either the full-length gene or a sufficiently large fragment of it to direct the synthesis of a protein product recognized by the primary antibodies. Once immunological screening of the genomic library is complete, it is necessary to determine which of the selected clones harbors the intact gene.

Fig. 4.14. Immunological screening of a genomic library (colony immunoblotting). Transformed cells are plated on a selective solid medium. 1. Cells from each colony are transferred onto a solid support (such as a nitrocellulose or nylon filter) to exactly match their arrangement on the agar plate. 2. The cells are lysed, and proteins are immobilized on the filter. 3. Primary antibodies that specifically bind to the target protein are added. 4. Unbound primary antibodies are washed away, and enzyme-linked secondary antibodies (e.g., conjugated to alkaline phosphatase) specific to the primary antibodies are applied. 5. Unbound secondary antibodies are washed off, and a colorless substrate is added, which yields a colored product upon Enzymatic hydrolysis. Hydrolysis occurs only in the presence of the secondary antibodies. 6. Colonies on the plate corresponding to the colored spots on the filter are selected and cultured. These may harbor recombinant DNA encoding a protein homologous to the one recognized by the primary antibodies.

Screening Based on Protein Activity

DNA hybridization and immunological methods enable the identification of numerous genes and their products. However, if the target gene encodes an enzyme that is not normally synthesized by the host cell, in situ plate assays can be used to identify clones carrying the gene. This strategy has been successfully employed to isolate genes for $\alpha$-amylase, endoglucanase, and $\beta$-galactosidase from various organisms. To do this, E. coli clones representing the genomic library of the organism of interest are plated on a nutrient medium containing a specific substrate. Following staining with a selective dye, cells capable of utilizing the substrate develop a distinctive coloration.

If the target gene encodes a product essential for the growth of a mutant host cell on minimal medium, the library can be constructed via the complementation of mutant cells by transformation. Cells that manage to grow on minimal medium in the absence of the required nutrient are guaranteed to carry a functional copy of the target gene delivered by the plasmid vector. Variants of this approach have been successfully utilized to isolate many important genes, including those responsible for antibiotic Biosynthesis and the formation of nitrogen-fixing ROOT nodules in certain plants.