Traditional cloning relies on recombinant DNA methods that begin with preparing a vector to receive an insert DNA by digesting each with restriction enzymes. The digested fragments are then spliced together by an enzyme called ligase, in a process known as ligation, to form a new vector capable of expressing a gene of interest. This may be the simplest and oldest technique for traditional cloning and laid the foundation for researchers to develop novel cloning methods such as TA cloning, TOPO cloning, PCR cloning, ligation-independent cloning, and gene assembly that exploit unique characteristics of other modifying enzymes.
A general workflow for traditional cloning includes the following steps (Figure 1):
Vectors used in traditional cloning methods are based on plasmids, which are double-stranded, circular DNAs that replicate inside bacteria independently of the genomic DNA. All cloning vectors based on plasmids contain a number of crucial elements, including a bacterial origin of replication to efficiently propagate within the bacterial host cell; single restriction enzyme site(s) or, more commonly, a multiple cloning site (MCS) that contains a number of restriction enzyme sites to allow ready addition of an insert of interest; and a marker (e.g., antibiotic resistance) to select for bacteria after successful uptake of the vector.
In some vectors, the MCS is located within a gene that serves as a marker and permits screening for clones into which the insert has been spliced successfully. For instance, the pUC18 vector expresses the lacZα gene encoding the alpha peptide of beta-galactosidase which, in combination with X-gal, allows color selection of bacterial colonies formed after cloning (learn more about blue/white selection in colony screening).
|Figure 2. Map of pUC18 with its MCS.|
The first step in preparing the vector for traditional cloning is to create an insertion site by restriction digestion. The choice of restriction enzymes depends upon the presence and location of their recognition sequences on the vector and the insert, and their compatibility for ligation. Vector restriction sites can be found on the vector map, or can be mapped using free online tools such as RestrictionMapper. The MCS, if available, is often the first choice for insertion, as the region is specifically designed for cloning.
After restriction digestion, dephosphorylation of the vector may be necessary to prevent self-ligation, especially if the resulting ends of vector digestion are compatible or blunt. During dephosphorylation, the enzyme alkaline phosphatase removes the 5′ phosphate groups at the ends. This prevents vector self-ligation because the enzyme ligase requires both a 5′ phosphate and a 3′ OH to join the two ends in recircularization of the vector (see Ligation). (App note: Dephosphorylation)
Dephosphorylation of the vector is important to reduce background and favor insertion of the desired fragment into the vector. Both self-ligated vector molecules and insert-carrying vector molecules can be taken up by the bacteria during transformation and will confer the same antibiotic resistance to those cells (see Colony screening and Figure 6). This will create a higher background of undesirable colonies if the vector is not dephosphorylated.
Blunting of the vector ends may be required, depending upon the restriction enzymes used. Purification of the desired fragments is also recommended for successful ligation.
The source of the insert for cloning may be genomic DNA, a portion of another plasmid, or a linear DNA fragment. Regardless of the type of source DNA, a common first step in preparation of the insert is to perform restriction digestion to generate compatible ends for subsequent splicing into the vector.
As with vector preparation, restriction enzymes that are suitable for cloning of the insert into the vector are selected. One of the most popular strategies is to perform double digests of both the insert and vector for directional cloning. In the following example (Figure 3A), two enzymes that generate non-compatible ends (EcoRI and KpnI) are used. Since vector and insert ends can join in only one orientation due to compatibility (EcoRI with EcoRI, KpnI with KpnI), this approach allows the insert to be cloned directionally. (App note: Directional cloning).
When performing double digestion, it is crucial that the reaction buffer and conditions are optimal for both enzymes; therefore, manufacturers’ recommendations for double digest reaction setups should be followed closely to ensure success. Some restriction enzymes are designed for complete digestion with multiple enzymes in a single buffer, enabling simplicity and time saving.
In instances where suitable restriction enzymes are not available, the DNA ends created by the chosen restriction enzymes may be blunted (or “polished”) for cloning. Blunting will alter the original sequences around the DNA ends; this could cause a frameshift in the gene translation region or disruption within a gene regulatory region. In addition, ligation of DNA with blunt ends is typically more challenging and less efficient than with cohesive (“sticky”) ends (Figure 3B).
In some instances, a single restriction enzyme may be chosen that cuts both the insert and vector DNA, generating complementary ends for ligation (Figure 3C). This method is commonly used in genomic DNA cloning.
In situations when it is not possible to use a single restriction enzyme, a pair of enzymes that have different recognition sequences but generate compatible overhangs can be considered as an alternative. For example, BamHI recognizes 5′-G↓GATCC-3′ and BglII recognizes 5′-A↓GATCC-3′; both generate 5′-GATC overhangs that can be joined in a ligation reaction. Note, however, that neither recognition site is restored after ligation in this case (Figure 3D).
Commonly used enzymes for generating blunt ends are the large (“Klenow”) fragment of DNA polymerase I, and T4 DNA polymerase. The choice of polymerase depends on whether the restriction enzyme generates a 3′ or 5′ overhang. In the case of 3′ overhangs (e.g., those generated by KpnI), T4 DNA polymerase is preferred because it has a stronger 3′ to 5′ exonuclease activity than does Klenow. In this scenario, the 3′ overhang is digested or “chewed back” by the T4 DNA Polymerase. For 5′ overhangs (e.g., those generated by EcoRI), either Klenow or T4 DNA Polymerase can be used to fill in the overhangs through their 5′ to 3′ polymerase activity. In a few instances, mung bean nuclease or S1 nuclease, added in excess, can be used to trim single-stranded DNA overhangs through their 5′ to 3′ exonuclease activities on single-stranded DNA (Figure 4).
|Figure 4. Enzymatic digestion to produce blunt ends from overhangs created by restriction digestion.|
After restriction digestion of the insert and the vector (and subsequent blunting and dephosphorylation, if performed), the desired fragments can be purified by running the samples on an agarose gel and excising the fragments of interest. Gel electrophoresis also removes enzymes and salts that were present in the digestion reactions. Gel purification kits are commercially available for efficient workflow, reliable results, and high yields. The kits are based on procedures that use a chaotropic reagent and heat to liquefy the gel, after which the fragments are purified using silica columns or magnetic beads. For gentle and efficient recovery of long DNA fragments (e.g., >10 kb), low melting gel, in combination with the enzyme agarase, which breaks down the agarose matrix, can be used. The traditional method for nucleic acid recovery from the solubilized gel is phenol/chloroform extraction, followed by ethanol precipitation of the fragments. The phenol/chloroform extraction may, however, result in lower yield and carryover of phenol that can affect downstream experiments. Extracted DNA should be highly pure for successful ligation. The simplest method to assess purity is to measure its absorbance: pure DNA has an A260/A280 ratio of >1.8 and an A260/A230 ratio of approximately 2.0.
Once the fragments of interest are obtained, a ligation reaction can be set up to join the insert and the vector. The most common enzyme used for ligation is T4 DNA ligase, which links DNA ends between 5′ phosphate and 3′ OH groups. The T4 DNA ligase reaction requires ATP, DTT, and Mg2+, which are generally supplied in the reaction buffer (Figure 5). To improve the outcome of ligation, a general recommendation is to set up multiple reactions with varying insert:vector molar ratios, typically in the range of 1:1 to 5:1. (Download the CloningBench app to access convenient tools and calculators, including the Vector to Insert Molar Ratios Calculator.) For less efficient ligations, as with DNA fragments with blunt ends, the addition of inert macromolecules like polyethylene glycol (PEG) is often recommended to increase the effective concentration of reaction components and thus improve the ligation efficiency.
|Figure 5. T4 DNA ligase reaction.|
Reaction temperatures may range from 14°C to 25°C (room temperature), and reaction times from 10 minutes to 16 hours (or overnight), depending on the type of DNA fragments and desired yields. In general, a higher reaction temperature requires less time but may produce a lower yield. Some commercially available ligation kits are designed to attain complete ligation in 15 minutes at room temperature. (App note: Ligation).
The ligated mixture may be used directly in transformation of chemically competent cells but may require purification prior to transformation of electrocompetent cells. If PEG was used in the ligation reaction, heat inactivation of the ligase is not recommended after the reaction, since this can reduce transformation efficiency.
Transformation is a naturally occurring process in which bacterial cells take up foreign DNA at a low frequency. In molecular biology applications, this process is enhanced and exploited to propagate plasmids inside bacteria that have been made “competent” (porous) for DNA uptake.
Competent cells are commercially available for efficient and reliable transformation. The most common approach to prepare bacteria to be competent for transformation is to treat log-phase bacterial cells with calcium chloride. When the chemically competent cells are mixed with the DNA from the ligation reaction and then heat-shocked at 42°C, some of the DNA is absorbed by the bacterial cells, where it begins to replicate.
Different strains of competent cells are available, and the choice is based on experimental goals and downstream applications. For instance, to perform “blue/white” screening, a bacterial strain with a lacZ mutation (lacZΔM15) must be chosen. If the experiment calls for digestion with methylation-sensitive restriction enzymes, the plasmid should be propagated in a dcm–/dam– bacterial strain. For protein expression, the strain should accommodate mRNA stability and translation, as well as high induction of the recombinant protein’s expression. (Learn more: Competent cell selection by applications).
In addition, transformation efficiency of the competent cells is an important consideration. Manufacturers provide the transformation efficiency of competent cells in colony-forming units per microgram of DNA (CFU/µg), generally ranging from 1 x 106 to 1 x 109 CFU/µg. In more difficult ligation and cloning strategies, choosing cells with the highest transformation efficiencies can greatly improve the likelihood of obtaining the desired clones.
Another method to transform bacterial cells is electroporation. In this technique, electrocompetent bacterial cells and ligated plasmids are treated with an electrical current that creates transient pores in the bacterial cell membrane for DNA uptake.
Transformed bacteria (after heat shock or electroporation) are then plated on an agar plate with an appropriate antibiotic, and screened (by blue-white screening or another method) for colonies that carry the desired plasmid with insert.
The transformation reaction contains a mix of cells with no vector, the vector with no insert, the insert alone, and the successfully ligated vector and insert. Bacteria without the vector lack the antibiotic resistance gene and will not grow, whereas bacteria transformed with the vector (with or without the insert) survive due to the expressed antibiotic resistance gene (Figure 6). Thus, the antibiotic resistance allows selection for uptake of an intact plasmid.
To identify whether the transformed colonies contain an insert, a number of methods can be employed, of which the most common are “blue/white” screening and positive selection. Blue/white screening relies on transforming a bacterial strain that expresses a mutant lacZ gene (lacZΔM15), which can be complemented with the alpha peptide of beta-galactosidase, encoded on the vector (alpha complementation). Transformed cells are plated on a growth medium that includes a transcriptional inducer for lacZ expression, IPTG, and a chromogenic substrate of LacZ, X-gal (5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside). In blue/white screening, LacZ will hydrolyze the X-gal, producing a blue dye and hence a blue colony. When a DNA insert disrupts the vector-encoded lacZα gene, no functional LacZ is formed, and transformed colonies are white (Figure 7).
Another popular screening approach is positive selection, whereby a gene lethal to the bacterial host is located in the MCS of the vector. Successful ligation of an insert into the lethal gene in the MCS prevents its expression, allowing only transformed cells with insert-carrying vectors to survive.
To more specifically identify or characterize the insert, transformed colonies must be further analyzed, as the results of blue/white screening and positive selection can only provide information about whether an insert is present or not. One basic approach is to perform restriction digestion of the vector extracted from the positive or white colonies and examine the resulting banding patterns from gel electrophoresis. Restriction enzymes must be carefully chosen and can be used to confirm the size and orientation of the insert.
The presence of the DNA insert can also be determined by a method called colony PCR, in which a small portion of the colonies is analyzed by PCR. This method requires PCR primers that are specific to the insert, to the flanking vector sequences, or both, to detect the insert. To determine the orientation of the insert, a set of primers that can detect the vector and the insert in a single reaction can be designed (Figure 8). (App note: Colony PCR).
The most definitive way to identify the insert is Sanger sequencing (also known as dideoxy sequencing). While this is a good way to confirm the presence and precise sequence of the insert, this approach may be time-consuming and cost-prohibitive, depending upon the number of colonies to be screened.
Once clones with the correct insert are identified, they are ready for downstream experiments.
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