Is PCR a DNA cloning technique?

Is PCR a DNA cloning technique?

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According to Genomes

PCR is

A technique that results in exponential amplification of a selected region of a DNA molecule [in test tube].

DNA cloning is

Insertion of a fragment of DNA into a cloning vector, and subsequent propagation of the recombinant DNA molecule in a host organism.

While, Molecular Biology of the Cell, Sixth Edition says in a summary:

DNA cloning (through the use of either cloning vectors or the polymerase chain reaction) in which portion of a genome (often an individual gene) is purified away from the remainder of the genome by repeatedly copying to generate many billions of identical molecules.

Here DNA cloning has been used to mean DNA replication in general.

Question: Which author's view is correct, or at least provides the most accepted definitions?

Short answer

The Oxford English Dictionary is quite clear on this. For the verb clone there are two meanings:

Biology To propagate (an organism or cell) as a clone.

Molecular Biology To make copies of (a DNA sequence or gene).

The latter definition clearly encompasses PCR.

History lesson

Most of the information below is taken from the Oxford English Dictionary.

The term clone has a long history, first as a noun and more recently as a verb. The evolution of the meaning and use of the word suggests that we shouldn't be too precious about this.

When I use a word,” Humpty Dumpty said, in rather a scornful tone, “it means just what I choose it to mean-neither more nor less.” “The question is,” said Alice, “whether you can make words mean so many different things.” “The question is,” said Humpty Dumpty, “which is to be master-that's all.

The word is based on a Greek word for twig and was originally used as a noun in the form clon.

1903 H. J. Webber in Science 16 Oct. 502/2 Clons… are groups of plants that are propagated by the use of any form of vegetative parts.

It soon gained the final e.

1905 C. L. Pollard in Science 21 July 88/1 I therefore suggest clone (plural clones) as the correct form of the word.

By 1930 it had become a verb…

1930 Jrnl. Ecol. 18 357 This is probably a record for the number of 'individuals' obtained at one time by cloning a herbaceous plant

… and, presumably because bacteria were considered to be plants at that time, it was used in microbiology too.

1929 Bibliographia Genetica 5 234 In Bacillus coli communis… a biotype was also found having lower motility than the remainder of the clone from which it came.

By the early 1960s it was extended to animals, probably as a direct result of Gurdon's experiments with Xenopus

1963 J. B. S. Haldane in G. Wolstenholme Man & his Future 352 Perhaps the first step will be the production of a clone from a single fertilized egg, as in Brave New World.

And finally the meaning was extended to encompass DNA molecules, thus molecular cloning.

1974 Proc. National Acad. Sci. U.S.A. 71 3459/1 ColE1 has been shown to serve as an effective molecular vehicle for cloning and amplifying specific regions of unrelated DNA.

(Extra points if you have read this far AND you know what ColE1 is.)

This final quotation seems to me to show that the confusion over what molecular biologists mean by cloning arose very early. If cloning means “to make copies of” then why is the word amplifying there? From the outset I think that molecular cloning carried the sense of purifying (by cutting and ligating). Or that's what I thought when I did it for the first time in 1980.


Despite my reliance on dictionary definitions above, and prompted by Tyto alba's discovery of a contradictory definition in a different OUP publication (see comment) here's what I really think:

Anyone who has ever obtained a bacterial colony containing a plasmid with the inserted fragment that they wanted announced "I have cloned the gene!" and meant that they had made a recombinant plasmid AND got it into cells. On the other hand, if they started with a PCR amplification then ran a gel to check before ligating and saw a fragment of the expected size they did not say "I have cloned the gene!" And finally, the successful cloner, when streaking out their transformant colony, will rarely have referred to this step as cloning.

Short answer
All of your sources are correct as they are not mutually exclusive. PCR is used to isolate and amplify DNA to yield small quantities of pure target product. Gene cloning can subsequently be applied to scale the production of the fragment up. PCR thus can be part of the DNA cloning process.

Cloning in general simply means duplicating. Cloning in Biology refers to

[Making] an exact copy of an animal or plant created in a laboratory from the animal's or plant's DNA

source: McMillan Dictionary

Gene cloning, as far as I understand refers specifically to the isolation of one single gene and inserting it into cells to amplify it:

The production of a lineage of cells all of which contain one kind of DNA fragment of interest derived from a population of many kinds of DNA fragments.

source: Northwestern University

PCR is a technique to amplify a specific strand of DNA that can be used to subsequently clone DNA into host cells (source: New England Biolabs). The cloning is done to upscale the quantity of DNA. PCR uses primers and can be used to isolate and amplify the DNA up to the scale it is easily clonable.

You may tell so. DNA amplification is a more generalized term for what you are indicating. Gene cloning with help of bacteria; popularly known as only Gene-cloning or Molecular cloning; is mainly used for gene or DNA amplification in vivo; and also PCR method (the most basic type) is also for DNA amplification, but in vitro, cell-free conditions.

But practically it may create some confusion; since the term "gene-cloning"/ "molecular cloning" popularly refer to that one done in living bacteria. (though the purpose or application is very same).

Reference: Wikipedia

Molecular Cloning Techniques

Molecular cloning or the creation of recombinant DNA is an essential process used in scientific research and discovery. With molecular cloning scientists can amplify and manipulate genes of interest and then insert them into plasmids for replication and protein expression. So how do scientists recombine DNA?

There are many methods that have been utilized over the years to move around pieces of DNA. Oftentimes several approaches will work for any specific cloning project however, it is likely that for any given project there is an ideal approach. This may be due to speed, cost, availability of starting materials or just personal preference. Check out our blog on choosing the right cloning method for your research project.

The following guide will highlight several of the most popular cloning methods used to create recombinant DNA.

What is Gene Cloning?

Gene cloning is a technique employed to locate and multiply a specific gene from the extracted genomic DNA of an organism through the construction of recombinant DNA. Genomic DNA contains thousands of different genes encoded for proteins. When DNA is extracted, it includes all possible genes it can bear. Gene cloning technique has enabled the detection of a specific gene from the total DNA. Therefore gene cloning serves as an important tool in molecular biology.

Making of a genomic library of an organism is essential in gene cloning if there is no clue about the location of the relevant gene in the DNA. A genomic library is made using the following steps.

Step 1: Extraction of the total DNA from an organism which contains the desired gene.

Step 2: Restriction digestion of the extracted DNA to produce small manageable fragments. This step is facilitated by restriction endonucleases.

Step 3: Selection of a suitable vector and opening the vector DNA using the same restriction endonucleases. Bacterial plasmids are commonly used as vectors to carry foreign DNA. Plasmids are small circles of DNA located within bacteria.

Step 4: Combining of the vector DNA and fragmented DNA to produce recombinant DNA molecule. This step is governed by DNA ligase.

Step 5: Transferring of recombinant DNA molecules into host bacteria. This step is known as transformation, and is done using a heat shock.

Step 5: Screening of transformed bacterial cells on a culture medium. A mixed population of transformed and nontransformed host cells is obtained at the end of the transformation process. As gene of interest includes only in transformed host cells. Hence, it is necessary to select transformed cells. The selection is made using selective media which contain antibiotics. Only the transformed cells grow on this screening medium enabling the selection.

Step 6: Growing of bacteria to produce a gene library. In this step, the transformed host cells are introduced into fresh culture media which provides optimum growth requirements. Total colonies on the culture plates represent the genomic library of that organism.

Step 7: The recombinant DNA molecule containing the gene of interest must be screened from thousands of cloned fragments of recombinant DNA. It can be accomplished by the use of probes which mark the specific gene or the specific protein results from that gene.

Once the interested gene containing the bacterial colony is identified from the total colonies, it is possible to make millions of copies of the recombinant plasmid that contains the gene.

Gene cloning is used in establishing gene libraries, producing special protein, vitamins, antibiotics, hormones, sequencing and mapping genomes of the organisms, making multiple copies of individuals DNA in forensics etc.

Figure_1: Gene Cloning

DNA polymerases

DNA polymerases are critical components in PCR, since they synthesize the new complementary strands from the single-stranded DNA templates. All DNA polymerases possess 5′→ 3′ polymerase activity, which is the incorporation of nucleotides to extend primers at their 3′ ends in the 5’ to 3’ direction (Figure 2).

In the early days of PCR, the Klenow fragment of DNA polymerase I from E. coli was used to generate the new daughter strands [3]. However, this E. coli enzyme is heat-sensitive and easily destroyed at the high denaturing temperatures that precede the annealing and extension steps. Thus, the enzyme needed to be replenished at the annealing step of each cycle throughout the process.

The discovery of thermostable DNA polymerases proved to be an important advancement, opening tremendous opportunities for the improvement of PCR methods by enabling longer-term stability of the reactions. One of the best-known thermostable DNA polymerases is Taq DNA polymerase, isolated from the thermophilic bacterial species Thermus aquaticus in 1976 [5,6]. In the first report in 1988 [7], researchers demonstrated Taq DNA polymerase’s retention of activity above 75°C, making continuous cycling without manual addition of fresh enzyme possible, and thus enabling workflow automation. Furthermore, compared to E. coli DNA polymerase, Taq DNA polymerase produced longer PCR amplicons with higher sensitivity, specificity, and yield. For all the aforementioned reasons, Taq DNA polymerase was named “Molecule of the Year” by the journal Science in 1989 [8].

Figure 2. DNA polymerase extending the 3′ end of a PCR primer in the 5′ to 3′ direction.

Although Taq DNA polymerase significantly improved PCR protocols, the enzyme still presented some drawbacks. Taq DNA polymerase is relatively unstable above 90°C during denaturation of DNA strands. This is especially problematic for DNA templates with high GC content and/or strong secondary structures that require higher temperatures for separation. The enzyme also lacks proofreading activity therefore, Taq DNA polymerase can misincorporate nucleotides during amplification. Where sequence accuracy is critical, PCR amplicons with errors are not desirable for cloning and sequencing. In addition, the error-prone nature of Taq DNA polymerase contributes to its inability to amplify fragments longer than 5 kb in general. To overcome such shortcomings, better-performing DNA polymerases are continually being developed to harness the power of PCR across a variety of biological applications (learn more about DNA polymerase characteristics).

Video: Fast PCR enzyme

Achieve 4x faster DNA synthesis, anneal primers at 60°C, and load samples directly onto gels after PCR, using Invitrogen Platinum II Taq Hot-Start DNA Polymerase.

PCR cloning strategies

PCR cloning is a method in which double-stranded DNA fragments amplified by PCR are ligated directly into a vector. PCR cloning offers some advantages over traditional cloning which relies on digesting double-stranded DNA inserts with restriction enzymes to create compatible ends, purifying and isolating sufficient amounts, and ligating into a similarly treated vector of choice (see insert preparation).

With PCR amplification, this cloning technique requires much less starting template materials which include cDNA, genomic DNA, or another insert-carrying plasmid (see subcloning basics). Furthermore, PCR cloning provides a simpler workflow by circumventing the requirement of suitably-located restriction sites and their compatibility between the vector and insert. Nevertheless, there are a number of considerations related to: PCR primers and amplification conditions, the cloning method of choice and the cloning vectors used, and, finally, confirmation of successful cloning and transformation.

With respect to PCR amplification of a sequence of interest, primers must be designed and PCR conditions (components and cycling) optimized for efficient and specific amplification of the template. Primer design tools are available to bioinformatically evaluate and select suitable target-specific primer sequences for amplification. Ligation requires that either the insert or vector has 5′-phosphorylated termini therefore, if the cloning vector lacks 5′-phosphorylated ends, 5′-phosphate groups must be added to the PCR primers during synthesis or by T4 polynucleotide kinase for successful ligation. For PCR optimization, reaction component concentrations, annealing temperatures, and template amounts are of importance.

TA cloning and blunt-end cloning represent two of the simplest PCR cloning methods. Their choice depends upon the nature of the vector and the type of PCR enzymes used in cloning. TA cloning employs a thermostable Taq DNA polymerase capable of amplifying short DNA sequences. This enzyme lacks 3′→ 5′ proofreading activity and features a terminal transferase activity that adds an extra deoxyadenine at the 3′ end of the amplicons (3′ dA). The resulting PCR products with 3′ dA overhangs are readily cloned into a linearized TA cloning vector containing complementary 3′ deoxythymine (3′ dT) overhangs (Figure 1). While relatively straightforward, the limitations of this method include the length of insert (up to 5 kb), the inability to clone inserts directionally, and the high error rate associated with Taq DNA polymerase.

Blunt-end cloning involves the ligation of an insert into a linearized vector where both DNA fragments lack overhangs. Blunt-end inserts can be produced using high-fidelity DNA polymerases with 3′→5′ exonuclease or proofreading activity. Their proofreading activity improves the sequence accuracy of the amplified products however, limitations include lower ligation efficiencies when inserting into blunt-end cloning vectors and the inability to clone directionally. Ligation efficiency can be improved by incubating the amplicons with a Taq DNA polymerase and dATP in a procedure called “3′ dA tailing” (incubate 20–30 minutes at 72°C), then purifying the 3′ dA-tailed products (Figure 1).

Figure 1. Common PCR cloning strategies.

To further simplify and streamline the cloning workflow, specialized vectors have been developed to place an insert into vector, for example, without using a ligase. One such class of vectors includes the Invitrogen TOPO cloning vectors which contain covalently linked DNA topoisomerase I that functions as both a restriction enzyme and a ligase (learn more about TOPO cloning technology). Compared to conventional PCR cloning vectors, these vectors result in shorter ligation reaction times (e.g., 5 minutes) and greater cloning efficiencies (e.g., >95% positive clones) and with a much simpler protocol. Furthermore, directional cloning of the PCR products can be achieved with a specially designed TOPO vector using a specific primer design.

Regardless of the cloning method choice, cloning efficiencies are significantly improved by purification of PCR amplicons prior to the ligation reaction. PCR clean-up helps remove salts, nucleotides, nonspecific amplicons, and primer-dimers. After ligation and transformation into the appropriate competent cells, the resulting colonies need to be screened carefully for the correct insert, as well as its proper frame and orientation for subsequent studies to analyze gene fusions and/or protein expression.

Polymerase Chain Reaction: Techniques and Variations

Read this article to learn about the techniques and variations of polymerase chain reaction with diagram.

Polymerase Chain Reaction :

The polymerase chain reaction (PCR) is a laboratory (in vitro) technique for generating large quantities of a specified DNA.

Obviously, PCR is a cell-free amplification technique for synthesizing multiple identical copies (billions) of any DMA of interest. Developed in 1984 by Karry Mullis PCR is now considered as a basic tool for the molecular biologist. As is a photocopier a basic requirement in an office, so is the PCR machine in a molecular biology laboratory!

Principle of PCR:

The double-stranded DNA of interest is denatured to separate into two individual strands. Each strand is then allowed to hybridize with a primer (renaturation). The primer-template duplex is used for DNA synthesis (the enzyme- DNA polymerase). These three steps— denaturation, renaturation and synthesis are repeated again and again to generate multiple forms of target DNA.

Technique of PCR:

The essential requirements for PCR are listed below:

1. A target DNA (100-35,000 bp in length).

2. Two primers (synthetic oligonucleotides of 17-30 nucleotides length) that are complementary to regions flanking the target DNA.

3. Four deoxyribonucleotides (dATP, dCTP, dCTP, dTTP).

4. A DNA polymerase that can withstand at a temperature upto 95° C (i.e., thermo-stable).

The reaction mixture contains the target DNA, two primers (in excess), a thermo-stable DNA polymerase (isolated from the bacterium Thermus aquaticus (i.e., Taq DNA polymerase) and four deoxyribonucleoties. The actual technique of PCR involves repeated cycles for amplification of target DNA.

Each cycle has three stages:

On raising the temperature to about 95° C for about one minute, the DNA gets denatured and the two strands separate.

2. Renaturation or annealing:

As the temperature of the mixture is slowly cooled to about 55° C, the primers base pair with the complementary regions flanking target DNA strands. This process is called renaturation or annealing. High concentration of primer ensures annealing between each DNA strand and the primer rather than the two strands of DNA.

The initiation of DNA synthesis occurs at 3′-hydroxyl end of each primer. The primers are extended by joining the bases complementary to DNA strands. The synthetic process in PCR is quite comparable to the DNA replication of the leading strand.

However, the temperature has to be kept optimal as required by the enzyme DNA polymerase. For Taq DNA polymerase, the optimum temperature is around 75° C (for E. coli DNA polymerase, it is around 37° C). The reaction can be stopped by raising the temperature (to about 95° C).

The 3 stages of PCR in relation to temperature and time are depicted in Fig. 8.1. Each cycle of PCR takes about 3-5 minutes. In the normal practice, the PCR is carried out in an automated machine.

As is evident from the Fig. 8.2 (cycle I), the new DNA strand joined to each primer is beyond the sequence that is complementary to the second primer. These new strands are referred to as long templates and they will be used in the second cycle.

For the second cycle of PCR, the DNA strands (original + newly synthesized long template) are denatured, annealed with primers and subjected to DNA synthesis. At the end of second round, long templates, and short templates (DNA strands with primer sequence at one end, and sequence complementary to the other end primer) are formed.

In the third cycle of PCR, the original DNA strands along with long and short templates are the starting materials. The technique of denaturation, renaturation and synthesis are repeated. This procedure is repeated again and again for each cycle. It is estimated that at the end of 32nd cycle of PCR, about a million-fold target DNA is synthesized (Table 8.1). The short templates possessing precisely the target DNA as double- stranded molecules accumulate.

Sources of DNA Polymerase:

In the original technique of PCR, Klenow fragment of E. coli DNA polymerase was used. This enzyme, gets denatured at higher temperature, therefore, fresh enzyme had to be added for each cycle. A breakthrough occurred with the introduction of Taq DNA polymerase from thermophilic bacterium, Thermus aquaticus. The Taq DNA polymerase is heat resistant hence it is not necessary to freshly add this enzyme for each cycle of PCR.

Key Factors for Optimal PCR:

Primers play a significant role in determining PCR. The primers (17-30 nucleotides) without secondary structure and without complementarity among themselves are ideal. The complementary primers can hybridize to form primer dimer and get amplified in PCR. This prevents the multiplication of target DNA.

As already described, Taq DNA polymerase is preferred as it can withstand high temperature. In the hot-start protocol, DNA polymerase is added after the heat denaturation step of the first cycle. This avoids the extension of the mismatched primers that usually occur at low temperature.

Taq polymerase lacks proof reading exonuclease (3′-5′) activity which might contribute to errors in the products of PCR. Some other thermo-stable DNA polymerases with proof-reading activity have been identified e.g., Tma DNA polymerase from Thermotoga maritama Pfu DNA polymerase from Pyrococcus furiosus.

In general, the shorter the sequence of target DNA, the better is the efficiency of PCR. However, in recent years, amplification of DNA fragments up to 10 kb has been reported. The sequence of target DNA is also important in PCR. Thus, CC-rich regions of DNA strand hinder PCR.

Promoters and inhibitors:

Addition of proteins such as bovine serum albumin (BSA) enhances PCR by protecting the enzyme DNA polymerase. Humic acids, frequently found in archeological samples of target DNA inhibit PCR.

Variations of PCR:

The basic technique of the PCR has been described. Being a versatile technique, PCR is modified as per the specific demands of the situation. Thus, there are many variations in the original PCR some of them are discussed, hereunder.

Nested PCR:

Sequence similarities between the target DNA and related DNA are very frequently seen. As a result of this, the primers may bind to both the DNAs and therefore even the undesired DNA also gets amplified in PCR. Use of nested primers increases the specificity of PCR, and selectively amplifies target DNA. Nested PCR is illustrated in Fig. 8.3. In the first cycle of PCR, the products are both from target DNA and undesired DNA. A second set of internal primers is now used. They will selectively bind to target DNA and amplification proceeds.

Inverse PCR:

In the inverse PCR, amplification of DNA of the unknown sequences is carried out from the known sequence (Fig. 8.4). The target DNA is cleaved with a restriction endonuclease which does not cut the known sequence but cuts the unknown sequence on either side. The DNA fragments so formed are inverted and get circularized (DNA ligase is employed as a sealing agent).

The circle containing the known sequences is now cut with another restriction enzyme. This cleaves only the known sequence. The target DNA so formed contains the known sequence at both the ends with target DNA at the middle. The PCR amplification can now be carried out. It may be noted that the primers are generated in the opposite direction to the normal, since the original sequence is inverted during circularization.

Anchored PCR:

In the anchored PCR, a small sequence of nucleotides can be attached (tagged) to the target DNA i.e., the DNA is anchored. This is particularly useful when the sequence surrounding the target DNA is not known. The anchor is frequently a poly G tail to which a poly C primer is used. The anchoring can also be done by the use of adaptors. As the adaptors possess a known sequence, the primer can be chosen.

Reverse Transcription PCR:

The PCR technique can also be employed for the amplification of RNA molecules in which case it is referred to as reverse transcription — PCR (RT-PCR). For this purpose, the RNA molecule (mRNA) must be first converted to complementary DNA (cDNA) by the enzyme reverse transcriptase. The cDNA then serves as the template for PCR. Different primers can be employed for the synthesis of first strand of cDNA. These include the use of random primers, oligo dT primer and a sequence specific primer (Fig. 8.5).

Asymmetric PCR:

PCR technique can also be used for the synthesis of single-stranded DNA molecules, particularly useful for DNA sequencing. In the asymmetric PCR, two primers in a ratio of 100: 1 are used. After 20-25 cycles of PCR, one primer is exhausted. The result is that in the next 5-10 PCR cycles, only single-stranded DNAs are generated.

Real-Time Quantitative PCR:

The quantification of PCR products in different cycles is not as simple as projected by theoretical considerations (Table 8.1). In practice, large variations occur. The most commonly used technique for measuring the quantity of PCR is by employing a fluorescence compound like eithidium bromide.

The principle is that the double-stranded DNA molecules bind to ethidium bromide which emit fluorescence that can be detected, and DNA quantified. The synthesis of genes by PCR and the role of PCR in site-directed mutagenesis are described elsewhere.

Random Amplified Polymorphic DNA (RAPD):

Normally, the objective of PCR is to generate defined fragments of DNA from highly specific primers. In the case of RAPD (pronounced as rapid), short oligonucleotide primers are arbitrarily selected to amplify a set of DNA fragments randomly distributed throughout the genome. This technique, random amplified polymorphic DNA is also known as arbitrarily primed PCR (AP-PCR).

The procedure of RAPD is comparable to the general technique of PCR. This method basically involves the use of a single primer at low stringency. A single short oligonucleotide (usually a 9-10 base primer) binds to many sites in the genome and the DNA fragments are amplified from them. The stringency of primer binding can be increased after a few PCR cycles. This allows the amplification of best mismatches.

RAPD can be carefully designed so that it finally yields genome- specific band patterns that are useful for comparative analysis. This is possible since genomic DNA from two different individuals often produces different amplified patterns by RAPD. Thus, a particular DNA fragment may be generated for one individual and not for the other, and this represents DNA polymorphism which can be used as a genetic marker.

RAPD is widely used by plant molecular biologists for the genetic identification of plant species. For this purpose, different combinations of nucleotides, most of them random oligonucleotide primers have been designed and are commercially available. As each random primer anneals to a different region of DNA, many different regions of loci on the DNA can be identified. RAPD is thus useful for the construction of genetic maps and as a method for genomic fingerprinting.

Limitations of RAPD:

The main problem of RAPD is associated with reproducibility. It is often difficult to obtain similar levels of primer binding in different experiments. It is therefore difficult to correlate results obtained by different research groups on RAPD.

Amplified Fragment Length Polymorphism (AFLP):

AFLP is a very Sensitive method for detecting polymorphism in the genome. It is based on the principle of restriction fragment length poly­morphism and RAPD. AFLP may be appropriately regarded as a diagnostic fingerprinting technique that detects genomic restriction fragments.

In the AFLP, PCR amplification rather than Southern blotting (mostly used in RFLP) is used for the detection of restriction fragments. It may be noted that AFLP is employed to detect the presence or absence of restriction fragments, and not the lengths of these fragments. This is the major difference between AFLP and RFLP. AFLP is very widely used in plant genetics.

It has not proved useful in the mapping of animal genomes, since this technique is mainly based on the presence of high rates of substitutional variations which are not found in animals. On the other hand, substitutional variations resulting in RFLPs are more common in plants. The basic principle of AFLP involves the amplification of subsets of RFLPs using PCR (Fig. 8.6).

A genomic DNA is isolated and digested simultaneously with two different restriction endonucleases — EcoRI with a 6 base pair recognition site and Msel with a 4 base pair recognition site. These two enzymes can cleave the DNA and result in small fragments (< 1 kb) which can be amplified by PCR. For this purpose the DNA fragments are ligated with EcoRI and Msel adaptors.

These common adaptor sequences (flanking genomic sequences) serve as primer binding sites on the restriction fragments. The DNA fragments can be amplified with AFLP primers each having only one selective nucleotide. These PCR products are diluted and used as templates for the selective amplification employing two new AFLP primers that have 2 or 3 selective nucleotides.

After the selective amplification by PCR, the DNA products are separated on a gel. The resultant DNA fingerprint is identified by autoradiography. AFLP fragments represent unique positions in the genomes, and hence can be used as landmarks to bridge the gaps between genetic and physical maps of genomes. In plants, AFLP is useful to generate high density maps, and to detect genomic clones.

Rapid Amplification of cDNA Ends (RACE):

As already described (See p. 115), reverse transcription, followed by PCR (RT-PCR) results in the amplification of RNA sequences in cDNA form. But the major limitation of RT-PCR is related to incomplete DNA sequences in cDNA. This problem is solved by using the technique rapid amplification of cDNA ends. RACE is depicted in Fig. 8.7, and briefly described below.

The target RNA is converted into a partial cDNA by extension of a DNA primer. This DNA primer was first annealed at an interval position of RNA, not too far from the 5′-end of the molecule. Now addition dATP (As) and terminal deoxynucleotidyl transferase extends the 3′-end of the cDNA.

This happens due to the addition of a series of as to the cDNA. These as series now act as the primer to anneal to the anchor primer. A second strand of DNA can be formed by extending the anchor primer. The double-stranded DNA is now ready for amplification by PCR. The above procedure described is called 5′- RACE, since it is carried out by amplification of the 5′-end of the starting RNA. Similar protocol can be used to carry out 3′-RACE when the 3′-end RNA sequence is desired.

Limitations of RACE:

Since a specific primer is used, the specificity of amplification of RACE may not be very high. Another disadvantage is that the reverse transcriptase may not fully reach the 5′-ends of RNA, and this limits the utility of RACE. In recent years, some modifications have been done to improve RACE.


Using the experimental procedures as outlined above, all four students in the semester-long laboratory course independently produced plasmids containing the GFP- His6-serpin chimera. This experimental series exposed the students to a number of modern techniques in molecular biology and allowed students to utilize these techniques in pursuit of a concrete project-based experimental goal. Students and faculty conferred as a group at the beginning and end of class periods. At the beginning of the semester, these meetings were used to check student calculations for the buffer preparation, to discuss time management during the remainder of the period, to comment on the day's protocols, and to introduce the theory underlying the protocols. Later in the semester, basic protocols (e.g. those supplied by vendors), computational tools, and websites were given to students and they were expected to design their experiments based solely on the vendor inserts this forced the students to become more comfortable with protocols and experimental time management. In addition, students were expected to bring their interpretation of their results to these meetings for discussion. Students became more independent in their work as the semester progressed.

This project continued in the following summer by two students and in the fall semester as an independent study. During these periods, the students have been able to express and purify the protein using a nickel affinity column to greater than 95% homogeneity from the bacterial expression system. The chimeric GFP-serpin product was demonstrated to have retained its serine protease inhibitory activity. As an additional educational benefit, the progress resulting from the work done in this laboratory sequence and summer session has generated the opportunity for student presentation at scientific conferences [ 16 ].

Furthermore, using the experimental procedures as outlined above, the four students participating in the independent project of Dr. Deibel were able to produce a set of plasmids containing the His6-N-lobe transferrin chimera. Students in this project conducted 6 h of research per week on days determined by their individual schedules. As a result, students needed to coordinate their research with one another, as the specific procedure they needed to run on any given day depended on what other students had completed the day before. During this project, students not only were exposed to “hands-on” research in modern molecular biological/biochemical techniques, but also were able to participate in a group research project.

National organizations (e.g. PKAL), and private and governmental funding agencies (e.g. Howard Hughes Medical Institute, National Science Foundation) have challenged the undergraduate science community to begin educating students in ways that better model the actual process of science. This laboratory series was designed to address these recommendations. Some of the potential advantages of a semester-long, research-based project are: 1) a closer resemblance to a professional research environment as might be encountered in graduate school 2) experience with experimental design 3) an understanding of how different techniques can be integrated to accomplish a larger goal 4) and a sense of personal involvement in a “real world” project that may lead to the advancement of knowledge and might be presented to the larger scientific community. Although this laboratory series was not strictly inquiry-based because a particular outcome was expected, the series does model the typical experiences involved in a research project, including the opportunities to trouble shoot steps that do not work. Such experiences establish technical and theoretical foundations on which students can then begin to do independent, inquiry-based work.

PKAL asserts that an important aspect of an undergraduate's success in a research environment is feeling the presence of a supportive community. To achieve this goal and because our students had different backgrounds in laboratory techniques, we encouraged them to rely heavily on each other, in addition to faculty. By being attentive to student successes and struggles, we were able to promote progressive independence in the laboratory. For instance, at the outset of the semester, protocols (e.g. for transformation) were rewritten and expanded by faculty from those supplied by the manufacturer in order to provide students with maximal instruction and explanation of theory. In some cases, presentations can be given using examples taken from educational biological textbooks to explain scientific principals involved in the protocol [ 17 – 21 ]. As the students came to understand the theory and which steps were critical, faculty provided abbreviated protocols and then monitored and questioned students closely during the laboratory to ensure they had a clear understanding of the protocol and how to carry it out. At the end of the semester, the students were provided with the protocols from the suppliers and expected to expand them into a more detailed one in their own notebooks. Consultation with faculty and other students was used to ensure that the students were interpreting the materials appropriately. This judicious withdrawal of support forced the students to pay attention to the overall goals of a protocol, as well as important details such as temperatures, incubation times, buffer compositions, and expected yields. At the end of the laboratory course, students were asked to give short detailed presentations on steps in the protocol, such as ligation or PCR, to reinforce their understanding of the principles involved in the performed laboratory procedures.

In addition to being representative of the professional research experience, the laboratory series modeled the importance of good laboratory record keeping, control experiments, and good experimental design. Although the timeline provided in Table I is representative of an ideal experimental series, the instructor must allow for experimental setbacks and be flexible in his/her guidance of the students over the course of the semester. Examples of experimental setbacks experienced by some of our students included failure to obtain purified DNA and failure to obtain colonies with the transformation following plasmid ligation. However, these occasional setbacks provided valuable educational opportunities. The sequential nature of the experimental series allowed for students to think about what might have gone wrong, adjust their experimental design, and then try the experiment again utilizing material from the previous step. Thus, both the instructor and the students should be aware of the amount of material (e.g. plasmid, PCR product) produced at each step while designing experiments, and save unused material in case problems arise in subsequent experiments. In some cases, materials such as plasmid preparations can be shared among students if one or more are unsuccessful in harvesting plasmid. In other cases, especially later in the semester when the students are more familiar with the experimental techniques, students can be encouraged to pursue independently whatever experiments are needed to complete the project.

This project-based learning format was well received by the students. In general, at the end of the semester, our students became very enthusiastic about completing their project and in several cases wanted to work on the project outside of normal laboratory hours. Student evaluations revealed that they considered the “hands on experience with biochemical techniques” and that “students were allowed to do much of the planning (and) experimental procedure” as positive aspects of the laboratory course. Additionally, all of the students indicated that this experience increased their interest in the subject. Finally, the plasmid constructs generated in the course have provided starting points for additional research projects for several students and likely will be featured in further presentations and publications. Thus, this experience appears to have provided a unique and valuable preparation for a career in experimental science.

The experimental sequence can be used to advance student or faculty research in a wide variety of situations. Such an approach may be especially beneficial for faculty at undergraduate institutions where the focus is on teaching and traditional research time is limited. Advances in DNA and protein technology will have an increasingly large societal impact. Accordingly, increased emphasis is currently being placed on the role of liberal arts colleges in introducing their students to relevant research projects. Fortuitously, the availability of new economical molecular biology kits and reagents is making the use of these techniques increasingly feasible in the undergraduate laboratory. The size of the laboratory section is certainly a consideration for such an open-ended approach. Our group, being unusually small, allowed for a great deal of individual attention and guidance. However, we are confident that such an approach could be used with a larger group of students. Peer-peer interactions were a significant resource, even among the group of four. Teaching assistants, which we did not have, could also provide a great deal of guidance and consultation and facilitate the use of this laboratory sequence with larger groups of students.

Because this series was successful, we feel there are several options for future Advanced Cell Physiology laboratories using this experimental sequence. One is to repeat the series with a different DNA amplification product as outlined in the preceding paragraph. As this procedure can be used with any sequence, a specific set of tagged genes such as those in a particular signal transduction pathway could be produced in different laboratory sections. Alternatively, a series of homologous tagged genes from of number of species could be produced. The recent completion of a number of genome projects has greatly increased the number of potential genetic targets. The generation of libraries of related tagged genes has a myriad of potential uses in further proteomic investigations of gene product function. Thus, the featured laboratory sequence could be used to substantially augment the research of undergraduate teaching faculty.

Overall cloning scheme to produce a GFP-His6-serpin construct. A, a map of the plasmid pGFPuv, which was purchased from Clontech. All plasmid maps were created by the shareware program MacPlasMap (v1.83). The GFPuv and ampicillin resistance genes are represented as black arrows. The arrows point in the direction of protein translation. The inducible lac promoter Plac controls GFPuv expression. The pUC origin of replication maintains a high copy number of the plasmid in the cell, facilitating plasmid purification. The multi-cloning sites (MCS) are shown in gray. The locations of the translational start and stop sites of the GFPuv gene, as well as the unique restriction sites SacI and EcoRI, are shown. The pGFPuv plasmid is digested with SacI and EcoRI and treated with the phosphatase CIAP, yielding a linearized DNA fragment that cannot relegate to form a circular plasmid. The sticky ends created by the restriction digestion of pGFPuv are used to ligate complementary sticky ends of the PCR insert containing the serpin cDNA. B, the PCR amplification product generated from plasmid serpin1B(A343K) containing the His6-serpin cDNA. The PCR primers are designed to introduce the complementary restriction sites to the ends of the His6-serpin DNA. These restriction sites must be unique in the PCR amplification product. Digestion of the PCR amplification product with SacI and EcoRI generates sticky ends suitable for ligation into the similarly linearized pGFPuv vector. C, the GFP-His6-serpin expression vector pMAJILK created by ligation of the linearized pGFPuv and the digested PCR amplification product. The complementary sticky ends created by the two restriction enzymes ensure proper directional ligation of the insert into the vector. The translation of the resulting 1.9-kb GFP-His6-serpin gene is under control of the inducible lac promoter. D, the protein product created by translation of the GFP-His6-serpin gene. This 75-kDa protein consists of a N-terminal 27-kDa GFPuv domain followed by a flexible linker domain that includes the His6 sequence. The C-terminal serpin domain is 45 kDa and contains the C-terminal protease cleavage site. The C-terminal protease cleavage site mandated that the GFP domain be added N-terminal of the serpin domain. The structures of the isolated GFP and serpin domains are also given. As shown, the figure is only approximately to scale. The flexible linker between the GFP and serpin domains consisting of glycine residues on either side of the His6 domain may promote accessibility of the epitope tag to solvent. Although the exact location of the domains in the recombinant protein is not known, the flexible linker appears to allow these domains to adopt positions that do not functionally interfere. GFP (#1EMA) and M. sexta serpin (#1SEK) files were downloaded from the Protein Data Bank and manipulated using Deep View-Swiss Pdb Viewer ( The protein is depicted in standard ribbon diagram format. The linker region between the two proteins is shown as a linear B-strand to demonstrate the potential three-dimensional relationship of the individual domains.

Primer design and PCR of the His6-serpin cDNA. A, overview of primer design for PCR amplification of the His6-serpin gene. Shown is the double-stranded 1.2-kb His6 serpin cDNA in plasmid serpin1B(A343K). The large arrow denotes the direction of transcription. Following heat separation of the two strands, two synthetic oligonucleotide primers are added to serve as templates for PCR amplification. These primers consist of a 3′ complementary and 5′ noncomplementary region. The 3′ complementary regions of the primers serve as the initiation points for primer elongation, while the 5′ noncomplementary regions allow for introduction of restrictions sites to the PCR product. The front primer binds to the noncoding strand near at the start of the His6-serpin gene, and the back primer binds to the coding strand at the end of the His6-serpin gene. B, design of the front DNA primer. Shown is the front primer bound to the noncoding strand of the His6-serpin gene in the plasmid serpin1B(A343K). Single-letter amino acid abbreviations for each codon are shown above the coding strand. The 5′ noncomplementary end of the primer consists of the SacI restriction site and a 6-base overhang of random sequence to allow efficient cleavage of the SacI site. As the SacI site in the plasmid pGFPuv lies just before the end of the GFPuv gene, DNA sequence coding for the last two amino acids in GFPuv was added subsequent to the restriction site to maintain the full-length GFPuv gene upon subcloning of the PCR product into pGFPuv. The bases of the noncomplementary region of the plasmid are denoted as N. A glycine residue was also introduced to allow conformational flexibility of the pGFPuv domain relative to the rest of the gene product. The cleavage site of the coding DNA strand upon digestion with SacI is shown. The complementary region of the primer consists of the start of the His6-serpin gene and included an alanine linker residue prior to the start of the serpin domain. Periods represent long DNA sequences extending from the sequence shown. The arrow indicates the direction of primer extension in a PCR. C, design of the back DNA primer. Shown is the back PCR primer bound to the coding strand of the terminus of the His6-serpin gene in the plasmid serpin1B(A343K). Single-letter amino acid abbreviations for each codon are shown above the coding strand. The 5′ noncomplementary end of the primer consists of the EcoRI restriction site and a 6-base overhang of random sequence to allow efficient cleavage of the EcoRI site. The bases of the noncomplementary region of the plasmid are denoted as N. A glycine residue was also introduced to allow conformational flexibility of the pGFPuv domain relative to the rest of the gene product. The cleavage site of the coding DNA strand upon digestion with EcoRI is also shown. An additional stop codon was introduced after the endogenous stop codon to ensure termination of the gene product. Periods represent long DNA sequences extending from the sequence shown. The arrow indicates the direction of primer extension in a PCR.

An 1.2% agarose E-gel (Invitrogen) of DNA samples generated throughout the laboratory course. All samples were diluted to 20 μl and loaded directly onto the E-gel. Following electrophoresis at 60 V for 45 min, the gel was placed on a ultraviolet transiluminator and the image captured using a CCD camera. Lane 1 contains the Directload wide-range DNA marker (Sigma, St. Louis, MO). The DNA fragment lengths of the marker are given adjacent to the gel. Lane 2 contains a DNA preparation of uncut pGFPuv showing multiple bands due to plasmid supercoiling. Lane 3 contains the pGFPuv preparation cut singly but EcoRI yielding a DNA fragment of 3.3 kb. Lane 4 contains the pGFPuv plasmid preparation digested by SacI. Lane 5 contains the double digest of pGFPuv by EcoRI and SacI, yielding a single visible fragment of ∼3.3 kb. Lane 6 contains the ∼1.2-kb PCR amplification of the His6-serpin product generated from plasmid serpin1B(A343K) and using PCR primers that introduced EcoRI and SacI restrictions site to the ends of the PCR products. Lane 7 contains the double digest of the PCR product yielding an ∼1.2-kb DNA fragment with sticky ends. Lane 8 contains the dilute ligation reaction prior to the addition of ligase containing digest pGFPuv and PCR product. The PCR product is not visible in the diluted sample due to its small size. Lane 9 contains the dilute ligation reaction solution following ligation. A number of faint higher-order molecular mass bands may be seen that are not present in lane 8, indicating that the ligation was successful. A transformation of this reaction yielded a small number of bacterial colonies. Lane 10 is a double digest of a plasmid preparation from a bacterial colony obtained following the ligation reaction. This digestion yielded a single 3.3-kb fragment as in lane 5, as well as a 1.2-kb fragment as in lane 6, indicating that the plasmid consisted of the pGFPuv expression vector and included the recombinant His6-serpin PCR insert. Lane 11 is a double digest of a plasmid preparation from a bacterial colony obtained following the ligation reaction. This digestion yielded a single 3.3-kb fragment as in lane 5, indicating that the plasmid from this colony is pGFPuv and lacks the recombinant serpin insert. Lane 12 contains Directload wide-range DNA marker (Sigma).

Chromatogram from an automated DNA fluorescent sequencing run. Shown is for the His6-N-lobe transferrin plasmid chromatagram produced by automated flourescent sequencing (Genegateway). The chromatogram is viewed using chromas, which can be downloaded free of charge at The automated read of the chromatagram is displayed.

Week Primary laboratory activity Pre-laboratory manipulations
1 Transformation of serpin cDNA into DH5α
2 Plasmid DNA isolation and student-designed diagnostic restriction digest Set up overnight culture on the day prior to lab
3 E-gel of cut vector and primer design
4 Primer design and primer ordering Reconstitute PCR primers
5 PCR of gene of interest
6 Use of E-gels to check PCR product restriction digestions Store PCR product at –20 °C following PCR
7 Restriction digestion of plasmid and vector. CIAP treatment of vector.
8 Ligation and transformation of ligation product
9 Plasmid preparation DNA from transformants diagnostic digestion of transformant plasmid DNA Set up overnight cultures of transformants
10 Agarose gel of diagnostic DNA digestion to identify recombinant product design and order sequencing primers
11 DNA sequencing Reconstitute sequencing primers
12 Interpretation of sequencing results Open sequence data files

In vitro Mutagenesis

PCR Mutagenesis of Circular DNA

PCR mutagenesis can also be used to amplify the entire plasmid containing the gene of interest. One straightforward method, termed ‘inverted’ or ‘counter’ PCR, uses back-to-back primers ( Figure 2B ) one PCR primer serves as the mutagenic oligonucleotide and the other oligonucleotide primes from the opposite strand, adjacent to the mutagenic primer. The PCR product is a full-length, linear plasmid which is then phosphorylated and ligated before transformation. This method can readily be used to make deletion mutants by creating a gap between the primers. Variants of this method include ‘recombinant circle’ PCR ( Figure 2C ) and ‘recombination’ PCR, both of which rely on recombination of linear plasmids. In these techniques, two inverse PCRs are performed with gapped primers at different sites. These two mutant plasmids are then recombined in vitro by mixing and annealing (recombinant circle PCR) or in vivo (recombination PCR). The gaps are then repaired by the bacterial DNA repair machinery.

Standard PCR experiment overview

The PCR is used to amplify a specific DNA fragment from a complex mixture of starting material called template DNA. The sample preparation and purification protocols depend on the starting material, including the sample matrix and accessibility of target DNA. Often, minimal DNA purification is needed. However, PCR does require knowledge of the DNA sequence information that flanks the DNA fragment to be amplified (called target DNA).

From a practical point of view, a PCR experiment is relatively straightforward and can be completed in a few hours. In general, a PCR reaction needs five key reagents:

DNA to be amplified : also called PCR template or template DNA. This DNA can be of any source, such as genomic DNA (gDNA), cDNA, and plasmid DNA.

DNA polymerase : all PCR reactions require a DNA polymerase that can work at high temperatures. Taq polymerase is a commonly used one, which can incorporate nucleotides at a rate of 60 bases/second at 70 °C and can amplify templates of up to 5 kb, making it suitable for standard PCR without special requirements. New generations of polymerases are being engineered to improve reaction performance. For example, some are engineered to be only activated at high temperatures to reduce non-specific amplification at the beginning of the reaction. Others incorporate a “proofreading” function, important, for example, when it is critical that the amplified sequence matches the template sequence exactly, such as during cloning.

Primers : DNA polymerases require a short sequence of nucleotides to indicate where they need to initiate amplification. In a PCR, these sequences are called primers and are short pieces of single-stranded DNA (approximately 15-30 bases). When designing a PCR experiment, the researcher determines the region of DNA to be amplified and designs a pair of primers, one on the forward strand and one on the reverse, that specifically flanks the target region. Primer design is a key component of a PCR experiment and should be done carefully. Primer sequences must be chosen to target the unique DNA of interest, avoiding the possibility of binding to a similar sequence. They should have similar melting temperatures because the annealing step occurs simultaneously for both strands. The melting temperature of a primer can be impacted by the percentage of bases that are guanine (G) or cytosine (C) compared to adenine (A) or thymine (T), with higher GC contents increasing melting temperatures. Adjusting primer lengths can help to compensate for this in matching a primer pair. It is also important to avoid sequences that will tend to form secondary structures or primer dimers, as this will reduce PCR efficiency. Many free online tools are available to aid in primer design.

Deoxynucleotide triphosphates (dNTPs): these serve as the building blocks to synthesize the new strands of DNA and include the four basic DNA nucleotides (dATP, dCTP, dGTP, and dTTP). dNTPs are usually added to the PCR reaction in equimolar amounts for optimal base incorporation.

PCR buffer: the PCR buffer ensures that optimal conditions are maintained throughout the PCR reaction. The major components of PCR buffers include magnesium chloride (MgCl2), tris-HCl and potassium chloride (KCl). MgCl2 serves as a cofactor for the DNA polymerase, while tris-HCl and KCl maintain a stable pH during the reaction.

The PCR reaction is carried out in a single tube by mixing the reagents mentioned above and placing the tube in a thermal cycler.

The PCR amplification consists of three defined sets of times and temperatures termed steps : denaturation, annealing, and extension (Figure 1).

Each of these steps, termed cycles, is repeated 30-40 times, , doubling the amount of DNA at each cycle and obtaining amplification (Figure 2).

Let's take a closer look at each step.

1. Denaturation

The first step of PCR, called denaturation, heats the template DNA up to 95 ° C for a few seconds, separating the two DNA strands as the hydrogen bonds between them are rapidly broken.

2. Annealing

The reaction mixture is then cooled for 30 seconds to 1 minute. Annealing temperatures are usually 50 - 65 ° C however, the exact optimal temperature depends on the primers' length and sequence. It must be carefully optimized with every new set of primers.

The two DNA strands could rejoin at this temperature, but most do not because the mixture contains a large excess of primers that bind, or anneal, to the template DNA at specific, complementary positions. Once the annealing step is completed, hydrogen bonds will form between the template DNA and the primers. At this point, the polymerase is ready to extend the DNA sequence.

3. Extension

The temperature is then raised to the ideal working temperature for the DNA polymerase present in the mixture, typically around 72 ° C, 74 ° C in the case of Taq.

The DNA polymerase attaches to one end of each primer and synthesizes new strands of DNA, complementary to the template DNA. Now we have four strands of DNA instead of the two that were present to start with.

The temperature is raised back to 94 ° C and the double-stranded DNA molecules – both the "original" molecules and the newly synthesized ones – denature again into single strands. This begins the second cycle of denaturation-annealing-extension. At the end of this second cycle, there are eight molecules of single-stranded DNA. By repeating the cycle 30 times, the double-stranded DNA molecules present at the beginning are converted into over 130 million new double-stranded molecules, each one a copy of the region of the starting molecule delineated by the annealing sites of the two primers.

To determine if amplification has been successful, PCR products may be visualized using gel electrophoresis, indicating amplicon presence/absence, size and approximate abundance. Depending on the application and the research question, this may be the endpoint of an experiment, for example, if determining whether a gene is present or not. Otherwise, the PCR product may just be the starting point for more complex downstream investigations such as sequencing and cloning.

Site-Directed Mutagenesis

Site-directed mutagenesis is a widely used procedure for the study of the structure and function of proteins by modifying the encoding DNA. By using this method, mutations can be created at any specific site in a gene whose wild-type sequence is already known. Many techniques are available for performing site-directed mutagenesis. A classic method for introducing mutations, either single base pairs or larger insertions, deletions, or substitutions into a DNA sequence, is the Kunkel method.

The first step in any site-directed mutagenesis method is to clone the gene of interest. For the Kunkel method, the cloned plasmid is then transformed into a dut ung mutant of Escherichia coli. This E. coli strain lacks dUTPase and uracil deglycosidase, which will ensure that the plasmid containing the gene of interest will be converted to DNA that lacks Ts and contains Us instead.

The next step is to design a primer that contains the region of the gene which you wish to mutate, along with the mutation you want to introduce. PCR can then be used with the mutated primers to create hybrid plasmids each plasmid will now contain one strand without the mutation and uracil bases, and another strand with the mutation and lacking uracil.

The final step is to isolate this hybrid plasmid and transform it into a different strain that does contain the uracil-DNA glycosylase (ung) gene. The uracil deglycosidase will destroy the strands that contain uracil, leaving only the strands with your mutation. When the bacteria replicate, the resulting plasmids will contain the mutation on both strands.

Experimental Procedure

Run PCR and purify the PCR product:

Run PCR to amplify your insert DNA. It is important to use a high fidelity taq polymerase to minimize mutations. The fidelity of the polymerase becomes more important the longer the expected PCR product is. You should select an annealing temperature based on the melting temperature (Tm) of the portion of the primer that hybridizes to the sequence to be amplified (the ORF in this case), not the Tm of the entire primer. If you are amplifying from a plasmid or simple template, there is very little chance for mis-priming, so you can use a pretty wide range of annealing temperatures, but you may need to increase your primer length and increase the Tm if you are trying to clone from genomic DNA, a cDNA library, or by RT-PCR.

Isolate your PCR product from the rest of the PCR reaction using a kit, such as the QIAquick PCR Purification Kit. The PCR product is now ready for restriction digestion.

Digest your DNA:

Set up restriction digests for your PCR product and recipient plasmid. Because you lose some DNA during the gel purification step, it is important to digest plenty of starting material. We recommend using your entire PCR reaction and 1μg of recipient plasmid. It is also critical that as much of the recipient plasmid as possible be cut with both enzymes, and therefore it is important that the digest goes at least 4 hours and as long as overnight.

If you are going to use only one restriction enzyme, or enzymes that have compatible overhangs or no overhangs after digestion, you will need to use a phosphatase to prevent re-circularization of the vector. You should treat your digested recipient vector with a phosphatase prior to the ligation step or prior to the gel purification step, depending on the phosphatase you choose. CIP (calf alkaline phosphatase) or SAP (shrimp alkaline phosphatase) are commonly used. Follow the manufacturer’s instructions.

Isolate your insert and vector by gel purification:

Run your digest DNA on an agarose gel and conduct a gel purification to isolate the DNA. When running a gel for purification purposes it is important to have nice crisp bands and to have space to cut out the bands. Because of this we recommend that you use a wide gel comb, run the gel on the slower side, and skip lanes between samples. In addition to a DNA ladder standard, it is also a good idea to run an uncut sample of each vector to help with troubleshooting if your digests don’t look as you expected.

When cloning by PCR, it is especially important to run the product on a gel. This allows you to visualize that your PCR product is the anticipated size and that your band is strong (indicating that the PCR reaction worked and that you have a sufficient amount of DNA).

Once you have cut out and purified your insert and vector bands away from the gel, it is important to determine the concentration of recovered DNA.

Ligate your insert into your vector:

Conduct a DNA Ligation to fuse your insert to your recipient plasmid.

We recommend around 100ng of total DNA in a standard ligation reaction. You ideally want a recipient plasmid to insert ratio of approximately 1:3. Since the number of base pairs for each varies, it is difficult to calculate this based on DNA concentration alone. One method is to conduct 2 ligations for each plasmid you are trying to create, with varying ratios of recipient plasmid to insert.

It is also important to set up negative controls in parallel. For instance, a ligation of the recipient plasmid DNA without any insert will tell you how much background you have of uncut or self-ligating recipient plasmid backbone.


Proceed with the transformation according to the manufacturer’s instructions for your competent cells.

For most standard cloning, you can transform 1-2μl of your ligation reaction into competent cells such as DH5alpha or TOP10. If using much less total DNA (<1ng) or if you are having trouble getting colonies, you might want to use higher competency cells. Additionally, if your final product is going to be very large (>10kb) you might want to use electro-competent cells instead of the more common chemically-competent cells.

The number of bacterial colonies resulting from your transformation will give you the first indication as to whether your transformation worked. Your recipient plasmid + insert plate should have significantly more colonies than the recipient plasmid alone plate. The recipient plasmid alone control will tell you your “background” level or more specifically it will tell you how many colonies you can expect on your recipient plasmid + insert plate that are not correct.

If you have a high number of colonies on your recipient plasmid alone plate, you can try ligating the recipient plasmid alone in the presence and absence of ligase. If the colonies are a result of uncut empty plasmid, you will still have colonies when you do not add ligase. If the colonies are a result of recipient plasmid self-ligation, you will see significantly more colonies when you add ligase.

If you do not see any colonies, you should conduct a positive control to ensure that your transformation worked. You could also try varying the amount of recipient plasmid to insert.

Isolate the Finished Plasmid:

Finally, you will need to pick individual bacterial colonies and check them for successful ligations. Pick 3-10 colonies depending on the number of background colonies on your control plate (the more background, the more colonies you will need to pick) and grow overnight cultures for DNA purification.

After purifying the DNA, conduct a diagnostic restriction digest of 100-300ng of your purified DNA with the enzymes you used for the cloning. Run your digest on an agarose gel. You should see two bands, one the size of your vector and one the size of your new insert.

Verify your Plasmid by Sequencing:

PCR based cloning carries a much higher risk for mutation than restriction enzyme based cloning. DNA replication by PCR has error rates that range from roughly 1 per 500bp to roughly 1 per 10 million bp depending on the polymerase used. Because of this, no matter which taq polymerase you use, it is important that you sequence the final product.