Molecular cloning is a laboratory method for creating numerous identical copies of a specific DNA sequence. This process can be likened to photocopying a single page from a vast library, allowing for its detailed study and use. The technique involves inserting a piece of DNA into a carrier molecule and introducing it into a host organism, which then replicates it. This capability to amplify a particular segment of DNA is important in many areas of biological and medical research.
The applications of molecular cloning are widespread, underpinning advancements in medicine and biotechnology. For example, it is the basis for the large-scale production of therapeutic proteins like insulin for treating diabetes and for the development of vaccines. In research, it allows scientists to investigate gene function, study protein interactions, and engineer organisms with new traits.
Preparation of the Vector and Insert
The initial step in molecular cloning requires the preparation of two primary DNA components: the “insert” and the “vector.” The insert is the specific DNA fragment of interest, which could be a gene, a regulatory element, or another sequence to be studied. The vector acts as a vehicle, a small, circular piece of DNA known as a plasmid, which carries the insert and facilitates its replication inside a host organism. Both components must be purified to remove contaminants like proteins, RNA, or solvents that could interfere with subsequent steps.
A common method to generate the insert is through the Polymerase Chain Reaction (PCR). PCR allows scientists to amplify a specific segment of DNA from a larger template, such as genomic DNA, creating millions of copies of the target sequence. The primers used in the PCR can be designed to add specific sequences to the ends of the amplified fragment, which are often recognition sites for restriction enzymes.
Once the insert is amplified and the vector plasmid is isolated, both are cut with restriction enzymes. These enzymes recognize and cleave DNA at specific, short sequences known as restriction sites. To ensure the insert is incorporated into the vector in the correct orientation, two different restriction enzymes are often used. This creates non-compatible ends on the vector and corresponding compatible ends on the insert, known as “sticky ends” because they have short, single-stranded overhangs that can base-pair with each other. After digestion, the desired DNA fragments are purified using agarose gel electrophoresis to remove unwanted DNA pieces.
Ligation of DNA Fragments
With the insert and vector prepared with compatible ends, the next stage is to join them together. This joining process is called ligation and is mediated by an enzyme known as DNA ligase, which functions as a molecular glue. The most commonly used enzyme for this purpose is T4 DNA ligase. This enzyme catalyzes the formation of a phosphodiester bond between the sugar-phosphate backbones of the DNA fragments.
The reaction is set up by mixing the purified insert DNA and the linearized vector DNA in a tube with the DNA ligase and a buffer containing ATP, which the enzyme requires for energy. The complementary sticky ends of the insert and vector anneal through hydrogen bonds, bringing the ends into close proximity. The T4 DNA ligase then acts on the nicks in the DNA backbone, sealing the gaps and permanently joining the two molecules.
This process results in the creation of a new, single, circular molecule called a recombinant plasmid. While T4 DNA ligase can join both sticky and blunt ends, ligation is generally more efficient with the complementary overhangs provided by sticky ends.
Transformation into Host Cells
After successfully creating recombinant DNA, the plasmids are introduced into living cells that will act as factories to produce many more copies. This process is known as transformation. The most common host organisms used in molecular cloning are laboratory strains of the bacterium Escherichia coli (E. coli). These strains are specifically selected for their inability to modify or degrade foreign DNA and their high efficiency in taking it up from the environment.
Before transformation, the E. coli cells must be made “competent,” meaning their cell walls are altered to become more permeable to DNA. Chemical transformation involves treating the bacteria with a solution of calcium chloride (CaCl2) and keeping them on ice. The positively charged calcium ions are thought to neutralize the negative charges on both the plasmid DNA and the bacterial cell membrane, reducing the electrostatic repulsion between them.
The plasmid DNA is then mixed with the competent cells and subjected to a “heat shock.” The mixture is moved from ice to a 42°C water bath for a short period, typically 30 to 45 seconds, before being rapidly returned to the ice. This sudden temperature change is believed to create transient pores in the cell membrane, allowing the plasmids to enter the bacterial cell. An alternative, more efficient but also more complex method is electroporation, which uses an electric pulse to create these temporary pores. Following the uptake, the cells are allowed a recovery period in a nutrient-rich medium before they are prepared for the next step.
Selection and Screening of Clones
Because the transformation process is not perfectly efficient and some cells may take up an empty vector without the insert, a two-part process of selection and screening is used to identify the correct bacteria. Selection is the first filter to eliminate bacteria that did not take up any plasmid. Cloning plasmids carry a selectable marker, a gene conferring resistance to a specific antibiotic. After recovery, the bacteria are spread onto an agar plate with this antibiotic, and only bacteria that have taken up a plasmid will survive to form colonies.
Screening then distinguishes between colonies with the desired recombinant plasmid and those with an empty one. A widely used technique is blue-white screening, which utilizes a vector containing the lacZα gene. When the DNA insert is successfully ligated, it disrupts the lacZα gene, rendering it non-functional. The bacteria are grown on a medium with X-gal, a substrate that turns blue when cleaved by the functional enzyme. Colonies with an empty vector appear blue, while colonies with the recombinant plasmid appear white.
Clone Verification and Propagation
After successful selection and screening, it is important to verify that the white colonies contain the correct plasmid with the insert properly integrated. A preliminary check can be done using colony PCR, but more definitive verification methods are typically employed. One common technique is to grow a small liquid culture from a single white colony and then isolate the plasmid DNA in a process called a miniprep.
This purified plasmid DNA can then be analyzed by restriction digest. The plasmid is cut with one or more restriction enzymes, and the resulting fragments are separated by size on an agarose gel. The pattern of DNA bands on the gel can confirm the presence and orientation of the insert by comparing the observed fragment sizes to the predicted sizes from the known plasmid and insert sequences.
For definitive verification, the isolated plasmid DNA is sent for DNA sequencing. Sanger sequencing is commonly used to read the exact nucleotide sequence of the insert and the junctions where it connects to the vector. This ensures that the insert’s sequence is correct, free of any mutations that might have been introduced during PCR, and is in the proper reading frame if it is intended for protein expression. Once a clone is fully verified, the bacteria containing it can be grown in a large-scale culture to produce a high yield of the recombinant plasmid for use in subsequent experiments.