DNA cloning means making identical copies (clones) of a DNA molecule using a DNA polymerase to replicate the DNA. Although it can be carried out in cell-free systems as well as in intact cells, the term is widely used to mean cloning DNA within intact cells, using a DNA polymerase naturally resident within the cells. The DNA sequences to be cloned must first be transferred into some suitable cells that can proliferate rapidly in culture. Cells that have correctly taken up the DNA of interest must be selected in some way, and allowed to proliferate in culture, resulting in a large increase in DNA copy number. After lysing the cells, the desired DNA sequences of interest are purified in some way.
Fractionating and purifying DNA by transforming bacterial cells with recombinant DNAs
The cells used in DNA cloning are typically well-studied bacterial cells, notably strains of Escherichia coli. They grow well in culture (and can quickly be expanded to very large numbers in large-volume containers), and they are amenable to genetic modification that maximizes the efficiency of the cloning process. Yeast cells are also used to clone very large DNA fragments.
The procedure initially involves treating the cells in some way so as to allow transfer of the DNA molecules that we wish to clone into the cells, a process known as transformation. In each case, the DNA to be cloned is covalently joined (ligated) to some vector DNA sequence that will help it replicate within the host cells, as detailed below.
The joining of DNA fragments to vector molecules results in the formation of an artificial recombinant DNA that may be linear in specialized cases (as in the case of cloning very large fragments in yeast). Usually, however, DNA cloning is carried out in bacteria where recombinant DNA molecules are circular (bacteria are more readily transformed by circular DNA).
The transformation process is selective: when foreign DNA does get into a cell, just a single DNA molecule is usually taken up by the cell. If a cell population is presented with a mixture of different foreign DNA fragments, therefore, different DNA fragments will be randomly allocated to different cells during transformation. That is, the population of cells serves as a kind of postal sorting office that can efficiently fractionate a complex mixture of DNA fragments (Figure 1).
Fig1. Transformation as a way of fractionating a complex sample of DNA fragments. The key point is that transformation is selective: when a cell is transformed it usually picks up a single DNA molecule from the environment, and so different fragments are taken up by different cells. Cell clones can form by repeated cell division from a single transformed cell and be propagated to produce a large number of cells with an identical foreign DNA sequence that can be purified after breaking the cells open. (Note: for clarity, the figure shows only the DNA sequences that are to be cloned—in practice they would be joined to a vector molecule.)
Amplification
Bacterial cloning systems offer the chance of making large quantities of a cloned DNA. That is, the inserted DNA is amplified to very high copy numbers as indicated in images at the right in Figure 1. That is possible for two reasons. First, a single bacterium containing a cloned DNA can rapidly divide, leading eventually to a huge number of identical bacterial cell clones, each with the same foreign DNA sequence. Second, some vector molecules can replicate within a bacterial cell to reach quite high copy numbers; if they have a foreign DNA sequence covalently linked to them, that too will be amplified within the cell (Figure 2). We consider some of the details below.
Fig2. Recombinant DNA may be amplified to high copy number within individual cells. Vectors have their own replication origin and can replicate within a bacterial cell independently of the host chromosome, often replicating much more frequently than the host-cell chromosome. For simplicity, the illustration here shows a very modest 3× amplification of the recombinant DNA, but some plasmids allow amplification to 100 copies or more in bacterial cells. A transformed bacterial cell can divide in culture to produce huge numbers of descendants that may each contain multiple copies of a recombinant DNA, resulting in a huge amplification of the starting recombinant DNA.
Vector molecules
Fragments of human DNA would not normally be able to replicate if transferred into bacterial cells or yeast cells. To replicate within cells, the DNA molecules need a suit able origin of replication, a DNA sequence that will initiate DNA replication in that cell type (molecules like this are known as replicons). A convenient solution is to take advantage of replication origins in DNA molecules that naturally replicate within the host cells.
For cloning in bacteria, extrachromosomal replicons are typically used that replicate independently of the bacterial chromosome. Two useful sources are plasmids (small, circular, double-stranded DNAs that can replicate to high copy numbers in some cases) and bacteriophages (bacterial viruses).
Plasmid vectors are popular because they are easy to work with, and they are versatile. Different plasmid vectors are suited to cloning fragments of different sizes (Table 1). Often, the object is simply to clone DNA fragments, but specialized plasmid vectors allow the insert DNA sequences to be expressed to produce RNA transcripts and proteins.
Table1. DIFFERENT PLASMID CLONING VECTORS TO ALLOW CLONING OF DNA FRAGMENTS OF DIFFERENT SIZES
To be useful as a cloning vector, the original plasmid, bacteriophage, or other replicon needs to be genetically modified so that we can efficiently join a foreign DNA to it (as described below) and so that transformed cells can easily be recognized. When cloning in bacteria, for example, the vector will have been genetically engineered to contain a gene that confers resistance to some antibiotic that the host cells are sensitive to. After transformation, the cells are grown on agar containing the antibiotic and untransformed cells die but transformed cells survive. Because some cells are transformed by naked vector DNA (lacking other DNA), screening systems are often also devised to ensure that cells with recombinant DNA can be identified.
Physical separation of clones
How can cells that have taken up different DNA fragments be separated from each other? The answer is to allow physically separated cell colonies to form. After transformation of bacterial cells, for example, aliquots of the cell mixture are spread over the surface of antibiotic-containing agar in Petri dishes (plating out); successfully transformed cells should grow and multiply and, if the plating density is optimal, they form well- separated cell colonies (Figure 3). Each colony consists of identical descendant cells (cell clones) that originate from a single transformed cell, and so the cell clones each contain the same single foreign DNA molecule.
Fig3. Picking well-separated bacterial colonies from a culture dish allows purification of cells containing a single type of recombinant DNA. Irrespective of whether a vector is capable of high-copy number amplification or not, any recombinant DNA can be amplified simply as a result of repeated division of the host cell. Growth occurs originally in a solid medium after the transformed cells are plated out; that is, spread out on a plate of agar containing nutrients and antibiotics (the vector is designed to contain a gene that confers resistance to the antibiotic; an additional selection system is often applied to ensure that the cells contain recombinant DNA, as described in the main text). During plating out, the cells will be physically dispersed to different parts of the agar surface in the culture dish, and individual surviving cells containing the antibiotic-resistance gene go through several rounds of cell division in situ to form visible colonies that may be well separated. An individual colony, consisting of identical cells with the same recombinant DNA molecule, can be picked and allowed to go through a second round of amplification by growth in liquid culture. For simplicity, the cloned DNA fragments are shown in the absence of the vector molecule.
An individual, well-separated cell colony can then be physically picked and used to start growth of a large culture of identical cells all containing the same foreign DNA molecule, resulting in very large amplification of a single DNA sequence of interest (Figure 3). Thereafter, the cloned foreign DNA can be purified from the bacterial cells.
Making recombinant DNA
To make recombinant DNA, each DNA fragment of interest needs to be covalently joined (ligated) by a DNA ligase to a vector DNA molecule. The resulting recombinant DNA molecules will subsequently be transported into suitable host cells, often bacterial or yeast cells. Before that is done, there is a need to prepare the DNA of interest and the vector DNA so that they can be joined efficiently, and there is a need to ensure that the recombinant DNAs are of optimal size.
To clone DNA in bacterial cells, we normally need to use relatively small DNA fragments. When DNA is isolated from the cells of complex organisms, the extremely long nuclear DNA molecules are fragmented by physical shearing forces to give a complex collection of still rather long fragments with heterogeneous ends. The long fragments need to be reduced to pieces of a much smaller, manageable size with uniform end sequences to facilitate ligation.
Recombinant DNA technology was first developed in the 1970s. The crucial break through was to exploit the ability of restriction endonucleases to cut the DNA at defined places. As a result, the DNA could be reduced to small, well-defined fragments with uniform end sequences that could be easily joined by a DNA ligase to similarly cut vector molecules (see Box1). Note that while most recombinant DNAs are circular, sometimes very large pieces of DNA are cloned in yeast cells and here the recombinant DNA is a linear DNA molecule called a yeast artificial chromosome (YAC), because it resembles a small yeast chromosome.
Box1. RESTRICTION ENDONUCLEASES: FROM BACTERIAL GUARDIANS TO GENETIC TOOLS
Plasmid copy number varies significantly: high-copy number plasmids may reach over 100 copies per cell, but other plasmids may be restricted to just 1–2 copies per cell. Different plasmids can coexist in a cell. A typical E. coli isolate, for example, might have three different small plasmids present in multiple copies and one large single-copy plasmid. Natural examples of bacterial plasmids include plasmids that carry the sex factor (F) and those that carry drug-resistance genes. Some plasmids sometimes insert their DNA into the bacterial chromosome (integration). Such plasmids, which can exist in two forms, extrachromosomal replicons or integrated plasmids, are known as episomes.
An example of standard DNA cloning in bacterial cells using a plasmid vector and a genetically modified host cell
In order to use natural plasmids (and phages) as vector molecules they need to be genetically modified in different ways. First, it is important to design the vector so that restriction fragments produced by cutting the sample DNA are inserted into a unique location (the cloning site) in the vector molecule. To allow cloning of different types of restriction fragments, the vector is genetically engineered to contain a 20–60 bp poly linker sequence with many different restriction sites. (As required, naturally occurring restriction sites in the vector are mutated and inactivated to ensure that the introduced restriction sites in the polylinker are unique—see top of Figure 4A for an example.)
Fig4. The high-copy number plasmid vector pUC19 and the basis of the lacZ color screen for recombinants. (A) Map of pUC19. The origin of replication (ori) enables more than 100 copies of pUC19 per host cell. The ampicillin-resistance gene (AmpR) permits selection for cells containing the vector molecule. A 54 bp polylinker (PL; uppercase letters outlined in red) has been inserted into lacZ′, a 5′ fragment of the lacZ (β-galactosidase) gene, and provides multiple unique cloning sites. lacI encodes a lac repressor protein (which represses transcription of lacZ′ until the inducer IPTG is added to the culture medium to bind to the repressor protein and inactivate it). (B) Basis of the lacZ complementation system and the blue/white color selection to identify recombinants. The host cell has been genetically modified to have a 5′ deletion in its lacZ gene so that it produces a protein lacking an N-terminal component of β-galactosidase. The vector’s 5′ lacZ′ sequence makes a polypeptide with the first 146 amino acids at the N-terminal end (α-fragment), but it is interrupted by 18 foreign amino acids as a result of translation of the inserted polylinker. The insertion is small and does not affect the activity of the α-fragment, and it can still complement the large C-terminal fragment of β-galactosidase (produced by the host cell’s lacZΔM15 gene) to make a functional β-galactosidase. In the β-galactosidase assay, a colorless substrate (Xgal, 5-bromo-4-chloro-indolyl-β-d galactopyranoside) is converted to 5-bromo 4-chloroindoxyl, which spontaneously dimerizes to give an insoluble, deep-blue pigment. Cloning of a large DNA fragment into the polylinker inactivates the α-fragment (either by introducing a frameshift or producing a much larger protein). Although the host cell continues to produce the large C-terminal fragment of β-galactosidase (not shown), it can no longer be complemented by an active α-fragment. Because no β-galactosidase can be produced, the cells will be colorless in the assay.
Vector selection
Another genetic modification is to have a marker gene in the vector for selecting only host cells that have been transformed by the vector. The plasmid vector pUC19, for example, contains a gene, AmpR, that confers resistance to the antibiotic ampicillin (Figure 4A), and the host E. coli cells used for cloning are ampicillin-sensitive. To screen for cells transformed by the vector, the transformed cells are plated out on an agar surface that contains the relevant antibiotic; the resulting colonies should be descendants of originally antibiotic-sensitive cells that have been transformed by the vector to become antibiotic-resistant.
Recombinant selection
Another type of genetic modification is to have a second marker gene in the vector for selecting only host cells that have been transformed by recombinant DNA. Sometimes the short polylinker is inserted into the coding sequence of the marker gene without changing the reading frame or disrupting the function of the marker gene. However, recombinants that have long inserts within the polylinker will disrupt the function of the marker gene (even if it does not change the reading frame, a large insert in the coding sequence will disrupt the function of the marker gene).
A widely used recombinant selection system derives from the E. coli lacZ gene and involves a color assay for β-galactosidase, the lacZ product. This enzyme can cleave the disaccharide lactose to give glucose plus galactose, but transcription of lacZ is normally repressed by a lac repressor protein encoded by the lacI sequence. Transcription of lacZ can, however, be induced by lactose, or a lactose analog such as isopropylthiogalactoside (IPTG), which binds to the repressor protein and inactivates it. IPTG is generally used as the inducer because it cannot be metabolized, and therefore its concentration does not change as the cells grow.
The lacZ marker assay is also designed to depend on complementation between different β-galactosidase fragments produced by the host cell and the vector. The E. coli host is a mutant in which the 5′ end of the lacZ gene has been deleted; it can produce a large C-terminal fragment of β-galactosidase, which by itself is nonfunctional. Vectors such as pUC19 (Figure 4A) have, however, been modified to contain a short 5′ component of the lacZ gene known as lacZ′ that when transcribed can produce a short N-terminal fragment of β-galactosidase (the α-fragment). The N-terminal α-fragment complements the host’s C-terminal β-galactosidase fragment to produce an active β-galactosidase that can be assayed by a reaction that gives a blue product (Figure 4B). However, because the cloning site is within the lacZ′ sequence of the vector, the lacZ′ sequence of recombinants is interrupted by a large insert and no β-galactosidase can be made.
The principle of DNA clone libraries and the applications and limitations of DNA cloning
Once DNA cloning was established it was soon used as a way of amplifying all DNA sequences from a starting source of DNA to make DNA libraries; that is, collections of DNA clones representing all types of DNA sequence in the starting DNA. DNA isolated from white blood cells, for example, provides a complex genomic DNA that can be cut into many pieces and attached to vector DNA molecules. The resulting mix of different recombinant DNA molecules is used to transform bacteria to produce very many different clones—a genomic DNA library. A good genomic DNA library would have so many different DNA clones that there was a good chance that the library included just about all the different DNA sequences in the genome.
An alternative was to make gene-centered DNA libraries. Until recently, it was imagined that the vast majority of human genes made proteins and an obvious starting point was mRNA. RNA cannot be cloned, however. Instead, DNA copies needed to be made using a specialized DNA polymerase, a reverse transcriptase that naturally copies a single-stranded RNA template to make a complementary DNA (cDNA) copy. Once the cDNA strand is made, the original RNA is destroyed by treatment with ribonuclease and the copied DNA strand is copied in turn to make a complementary DNA, thereby making double-stranded cDNA. Total double-stranded cDNA isolated from cells could then be used to make a cDNA library. But because different genes are expressed in different cell types, the range of DNA clones in a cDNA library could vary according to whether the cDNA originated from white blood cells or brain cells, and so on.
Once genomic and cDNA libraries were made from human cells (and the cells of model organisms) they could be screened using previously isolated DNA clones (or syn thetic oligonucleotides) as probes to identify related DNA clones. We go through the details of how this is done in Chapter 7, where we also describe specialized large-insert DNA cloning systems that were first widely used in genome projects.
DNA cloning started a revolution in genetics. It prepared the way for obtaining panels of DNA clones representing all the sequences in the genome of organisms, making genome projects possible, and comprehensive collections of expressed sequences in different types of cell. To allow expression of coding DNAs, cDNA clones were sub cloned into specialized plasmid vectors to produce large amounts of purified proteins that could be used for various purposes, including therapeutic purposes or for raising antibodies to track expression of that protein in cells and tissues. By cutting and pasting sequence components derived from different genes it became possible to create hybrid genes that have had different uses, including the production of different types of artificial antibody. By optimizing techniques for transferring gene/cDNA clones into human cells and expressing them within cells, a new field of gene therapy became possible.
There is a downside: cloning DNA in cells is laborious and time-consuming. It is therefore not suited to diagnostic testing and screening work (where parallel amplifications of one or more DNA sequences need to be carried out rapidly in multiple different DNA samples). That required a new technology, the polymerase chain reaction (PCR).
Expression cloning in bacterial cells as a way of making large amounts of a desired protein
The cloning systems described so far in this section are designed simply to amplify the target DNA to obtain sufficient quantities for structural and functional studies. However, in many circumstances it is also useful to be able to express the introduced gene in some way. In expression cloning, appropriate signals need to be provided alongside the introduced gene to enable the gene to be expressed in the transformed host cell.
Depending on the type of expression product required, and the purpose of the expression, many different expression cloning systems can be used. Sometimes an RNA product is sufficient, but in many cases the object is to produce a protein. In some cases, all that is required is to analyze gene expression, for which low expression levels are accept able; in others, high expression is needed to retrieve, for example, large quantities of a specific protein. It is common to express the product in well-studied cells, but sometimes it may be sufficient to express the product in vitro.
Cloning of eukaryotic cDNA in an expression vector is often required for the pro duction of proteins in large quantities for research purposes such as structural studies, or for biotechnology purposes, or as medically relevant compounds such as therapeutic proteins. Usually, a cDNA providing the genetic information specifying the protein sequences is inserted into a vector along with separate expression signals such as suit ably strong promoters and other regulatory elements. Because the expression system is based on recombinant DNA, the resulting proteins are sometimes described, rather inaccurately, as recombinant proteins.
Bacterial cells, notably the well-studied Escherichia coli, have been widely used in expression cloning: they grow rapidly and can be expanded easily in culture to very large culture volumes. But the production of very large amounts of a nonendogenous protein can, however, be detrimental to the growth of the host cell, and can sometimes be toxic. An inducible promoter is often used, therefore, so that expression can be delayed until the transformed cells have been identified and grown in bulk. For example, the pET series of vectors contain a bacteriophage T7 promoter that is not recognized by the endogenous E. coli RNA polymerase (Figure 5). Such vectors are used to transform a genetically modified E. coli strain that contains a T7 RNA polymerase gene regulated by a lac promoter. The transformed cells can be selected and grown up in large quantities without expression of the foreign gene. Addition of the β-galactosidase inducer IPTG activates the lac promoter and expression of the adjacent foreign gene, and the cells can be harvested shortly afterward.
Fig5. Inducible bacterial expression vectors. The pET-3 series of plasmid vectors contain a bacteriophage T7 promoter. They are used with a strain of E. coli that has been genetically modified to contain a gene for the phage T7 RNA polymerase under control of an inducible lac promoter. After transformation by a pET-3 recombinant DNA, the E. coli cells are allowed to grow to give large numbers of cells in culture. At a desired stage, the lactose analog isopropylthiogalactoside (IPTG) is added to induce expression of the host’s T7 RNA polymerase gene. The induced T7 RNA polymerase specifically binds to the T7 promoter on the recombinant DNA to give high-level expression of the insert. AmpR, ampicillin-resistance gene; ori, origin of replication.
Bacteria have many advantages for expressing heterologous (foreign) proteins, but they also have limitations. Many eukaryotic proteins are modified by the addition of phosphate, lipid, or sugar groups after translation, and these modifications are often essential for the biological function of the protein. Bacterial cells lack the enzymes needed for eukaryotic post-translational processing, however, and so eukaryotic proteins artificially produced in bacterial cells often become unstable, or show limited or no biological activity. That has prompted the use of animal and mammalian cell systems for expressing many human and mammalian genes. We cover aspects of this when we consider genetic manipulation of human and animal cells in Chapter 8.
The need for fusion proteins and affinity tags
The large size of many eukaryotic proteins, notably mammalian proteins, also poses difficulties for synthesizing them in E. coli (where average protein sizes are only about 300 amino acids). Overexpression of large proteins leads to the production of insoluble aggregates of misfolded protein (inclusion bodies). The inclusion bodies can easily be purified, but it is difficult to then solubilize the protein and achieve efficient refolding in vitro.
Efforts to increase yield and solubility have often involved the production of fusion proteins. For example, the vector can be modified so that immediately adjacent to the cloning site it contains a cDNA sequence for all or part of an endogenous protein. Recombinants will therefore express the desired protein fused to an endogenous protein sequence. Many modern protein expression vectors are modified to contain a coding DNA sequence for a specific peptide or protein that is easily purified by affinity chromatography. In such cases the recombinant DNAs are expressed to give the desired protein but with a short peptide or protein tag attached that is known as an affinity tag because it is attached to assist purification of the recombinant protein by affinity chromatography.
Two favorite systems that allow affinity purification of expressed proteins are based on GST–glutathione affinity and polyhistidine–nickel ion affinity. Glutathione-S transferase (GST) is a small protein with a very high affinity for its substrate glutathione. The expression cloning vector positions the target DNA just after a gene encoding GST so that a GST-fusion protein is produced in the transformed host cell (Figure 6). This fusion protein can be purified by selective binding to a column containing glutathione. Alternatively, an affinity tag of six consecutive histidine residues can be attached to a protein. The side chains of the (His)6 tag bind selectively and strongly to nickel ions, assisting purification by affinity chromatography using a nickel–nitrilotriacetic acid matrix.
Fig6. Fusion protein vectors. The pGEX-4T series of vectors have a tac promoter (Ptac, a hybrid promoter with elements of the trp and lac promoters). Downstream gene expression is normally repressed by the repressor protein encoded by the lacIq gene but is inducible by the lactose analog IPTG. Immediately downstream of the tac promoter is a gene for the affinity tag glutathione S-transferase (GST) followed by a multiple cloning site (MCS). The object is to clone a target coding sequence for a protein into the MCS so that a fusion protein is produced, with an N-terminal GST sequence fused to the protein encoded by the target cDNA. Three alternative vectors, pGEX-4T-1, pGEX-4T-2, and pGEX-4T-3, have slightly different MCS sequences that differ in the translational reading frame (see from proline codon onwards in the sequences shown in the top half of figure). By using all three alternative vectors, cloned inserts can be expressed in each of the three amino acid reading frames so that in at least one of the three cases, the target DNA should be in frame with the GST sequence. The expressed fusion protein can be purified easily on a glutathione affinity purification column such as glutathione sepharose 4B. Because the MCS is engineered to contain a thrombin cleavage site, the desired protein can be purified after cleavage at the thrombin cleavage site. AmpR, ampicillin-resistance gene; ori, origin of replication; STOP, termination codon.
Phage display
Phage display involves inserting a coding DNA into a bacteriophage vector to produce a recombinant DNA that is transferred into bacteria, leading to the production of recombinant phage. The recombinants that express the foreign DNA give a protein that is dis played on the surface of a phage particle.
Because the cloning sites in the phage vector are designed to lie within a gene encoding a phage coat protein, expression results in a fusion protein that is incorporated into the phage’s protein coat, so that it is displayed on the surface of the phage (but it does not affect the phage’s ability to infect cells). If an antibody is available for the expressed protein, phage displaying the protein can be selected by preferential binding to the antibody: affinity purification of virus particles bearing such a protein can be achieved from a 108-fold excess of phage not displaying the protein, using even minute quantities of the relevant antibody (Figure 7).
Fig7. Phage display. (A) A heterogeneous mixture of target cDNAs is cloned into a bacteriophage vector, such as one based on phage M13 or f1, in order to express foreign proteins on the phage surface. Here, DNA is inserted into a cloning site at the extreme N-terminal sequence of the gene for protein III from phage f1 that makes one of the phage coat proteins. The recombinants are allowed to transform host E. coli cells, whereupon phage DNA replicates, and phage particles are assembled, extruded from the host cell, and harvested. (B) The mixture of recombinant phage is known as a phage expression library. Recombinants with in-frame inserts may often be expressed to give a fusion protein in which the N-terminal component consists of a protein sequence encoded by the inserted DNA. An antibody specific for one of the inserted proteins will bind specifically to just the phage that displays that protein. If the antibody bears an affinity tag such as biotin, it will selectively bind to its partner streptavidin, allowing purification of the labeled phage.
Phage display is a very versatile system. It is used in protein engineering to select for desired variants from a library of mutants. It has also proved a powerful alternative source of constructing antibodies, bypassing normal immunization techniques and even hybridoma technology. Phage libraries can also be used to identify proteins that interact with a specific protein. In the same way in which antibodies can be used in affinity screening, a known protein (or any other molecule to which a protein can bind) can be used as a bait to select phages that display any other proteins that bind to the bait protein.
