I am trying to do a PCR using a plasmid as the template, pretty basic stuff. But the problem is that my forward primer just happens to bind to another location on the plasmid in the reverse orientation, in between the proper forward binding site and the reverse binding site. This isn't an example of non-specific binding, the sequences are perfect because the plasmid contains self-complimentary regions and my forward primer just happens to land right on it.
Here is a diagram:
Simply optimizing the melting point or magnesium concentrations probably won't help, because the sequences are the same. I'm doing this cloning to insert a reporter gene into the vector, and I need to place it very precisely, so I don't know if I can order a new set of primers, the insert needs to fit there.
The bad PCR product is about 500 bases shorter than the good product, but I only see the bad band on gel when I run a normal PCR. If I reduce the amount of forward primer and increase the reverse oligo, I can see the good band, but the bad band is still there and the total yield is much lower.
So my idea right now is to do an asymmetric PCR using the reverse primer only, then add forward primer to create the double-stranded DNA, then separate the good and bad bands using gel electrophoresis. But I don't have a clue how many cycles I should run each step, or how much forward primer to add, etc. Additionally, asymmetric PCR has low yield due to arithmetic amplification, but once I add the forward primers it can begin symmetric PCR and geometrically amplify both bands, but I think the shorter bad band wins out.
Does anyone here have experience with this sort of complex PCR, or suggestions to improve my yield?
From experience, your present problem will not completely go away no matter what you do. The idea for asymmetric PCR is not a bad idea. However, you will only get linear amplification of the desired product from the initial single primer, so you will not "see" any product even after 30-40 cycles. This first single primer PCR might also cause downstream problems with amplification of undesired products made during the second PCR (junk products).
I agree with comment #1, but then neatly cut the "visible" "good" band out of the gel. Suspend (crush) the agarose band in sterile TE buffer, and then use an aliquot of that as template for a secondary PCR (use equal primers - maybe 30 cycles). This should provide more than enough "good" starting template.
Alternately, you could design a new forward primer that does not have a bad secondary location on your plasmid.
Polymerase Chain Reaction (PCR) | Biotechnology
The below mentioned article provides a Beginner’s Guide to Polymerase Chain Reaction (PCR). After reading this article you will learn about: 1. History of the PCR 2. The Principle of Polymerase Chain Reaction 3. Requirements of PCR 4. The PCR Reaction Cycle 5. Analysis of PCR Products 6. Applications 7. Precautions and Drawbacks 8. Modifications.
- History of the PCR
- The Principle of Polymerase Chain Reaction
- Requirements of PCR
- The PCR Reaction Cycle
- Analysis of PCR Products
- Applications of PCR
- Precautions and Drawbacks
1. History of the PCR:
The idea for PCR is credited to Kary-Mullis who was a research scientist in 1980s at a Cali­fornia biotechnology company called Cetus. Mullis, and five other researchers in Human Genetics Department at Cetus, demonstrated that oligonucleotide primers could be used to specifically amplify defined segments of ge­nomic DNA (or cDNA). Mullis was co-winner of 1993 Nobel Prize in Chemistry.
2. The Principle of Polymerase Chain Reaction:
Polymerase chain reaction (PCR) is a primer mediated enzymatic amplification of specifi­cally cloned or genomic DNA sequences. In this process we take the DNA with a target se­quence which we want to amplify, denature it by increasing the temperature and then use a sequence specific primer for the amplification of our target sequence by the help of a ther­mos-table DNA polymerase.
In this technique we try to reproduce an artificial environment under in vitro conditions in which the target DNA sequence undergoes multiple rounds of replication cycle to produce an enormous cop­ies of our target gene.
The process PCR has three fundamental steps:
Double-stranded DNA template denaturation by increasing the tempera­ture to 94 – 98°C for 30 seconds for 2 mi­nutes.
Annealing of two oligonucleotide primers to the single-stranded tem­plate by lowering the temperature to 50 – 65°C.
Enzymatic extension of primers to produce copies that can serve as tem­plates in subsequent cycles.
3. Requirements of PCR:
(a) DNA Template:
The original DNA mol­ecule that is to be copied is called the DNA template and the segment of it that will actually be amplified is known as the tar­get sequence. A trace amount of the DNA template is sufficient. This can be obtained by any one of the DNA isolation techniques discussed before.
(b) PCR Primers:
Two PCR primers are needed to initiate DNA synthesis. These are short pieces of single-stranded DNA that match the sequences at either end of the target DNA segment. PCR primers are made by chemical synthesis of DNA.
There are several computer programs available to suggest suitable primers for the process of PCR, and some of the general guide­lines are listed below:
Shorter primers have a tendency to go and anneal to the non-target sequence of the DNA template. This will result in production of DNA copies of having non-target sequence. The greater the complexity of the template DNA, the more likely this is to happen.
Thus, a Short primer may offer sufficient specificity when amplifying using a simple template such as a small plasmid, but a long primer may be re­quired when using eukaryotic genomic DNA as template. In practice, 20-30 nucleotides is generally satisfactory.
Primers do not need to match the template completely, although the 3′ end of the primer should be correctly base-paired to the template, otherwise the polymerase will not be able to extend it. It is often beneficial to have C or G as the 3′ terminal nucleotide. This makes the binding of the 3′ end of the primer to the tem­plate more stable than it would be with A or T at the 3′ end.
3. Melting Temperature:
The temperatures at which the two primers can associate with the template should be rela­tively similar to ensure that they both bind at about the same time as temperatures are be­ing lowered during annealing. The similarity of melting temperatures is likely to mean that the primers have a similar nucleotide compo­sition.
4. Internal Secondary Structure:
This should be avoided in order to prevent the primer to fold back on itself and not be avail­able to bind to the template.
5. Primer-Primer Annealing:
It is also important to avoid the two primers being able to anneal to each other. Extension by DNA polymerase of two self-annealed prim­ers leads to formation of a primer dimer.
(c) Thermo-Stable DNA Polymerase:
The enzyme DNA polymerase is needed to manufacture the DNA copies. The Klenow fragment was the first DNA polymerase enzyme used in PCR. The Klenow frag­ment is a large protein fragment produced when DNA polymerase I from E. coli is enzymatically cleaved by the protease subtilisin.
After enzymatic modification it retains the 5′-3′ polymerase activity and the 3′ → 5′ exonuclease activity for re­moval of pre-coding nucleotides and proof­reading, but loses its 5′ → 3′ exonuclease activity.
Klenow fragment failed to play a successful role as a polymerase enzyme for lacking a stability at high temperature. As we know that the PCR procedure involves several temperature steps, in this situa­tion we had to replenish the Klenow frag­ment during each cycle.
To solve this is­sued heat resistant DNA polymerase was required. This came originally from heat resistant bacteria living in hot springs at temperatures up to 90°C. Today Taq poly­merase from Thermusaquaticus is the most widely used PCR DNA polymerase enzyme. It is generally produced by expres­sion of the gene in E. coli.
The thermo-sta­bility of the Taq enzyme helps in its puri­fication after expression in E. coli, since- contaminating E. coli proteins can be in­activated by heating. The enzyme has 5′- 3′ DNA polymerase and 5′-3′ exonuclease activities. It will polymerize about 50-60 nucleotides per second. However, the en­zyme has a number of properties that may be disadvantageous.
1. Taq Polymerase has No Proof-Reading (3′-5′ exonuclease) Activity:
Consequently about one nucleotide in 10 4 in­corporated is incorrect and the individual prod­ucts of PCR will be a heterogeneous popula­tion.
2. Taq Polymerase has Relatively Low Processivity:
This means that it is likely to dissociate from the template before it has synthesized a long piece of DNA.
3. Taq Polymerase is not Fully Heat Stable:
It has a half-life of about 40 min at 95°C, which means there will be significant loss of activity over the 30 or so cycles used in a typical PCR experiment. It may, therefore, be necessary to add more enzyme during the course of an ex­periment.
4. Taq Polymerase Incorporates an Ex­tra a Residue:
This is incorporated on the 3′ end of the mole­cule synthesized, and is not template encoded. A number of polymerases are available from other Thermus species. These include Tfl and Tth enzymes from Thermusflavus and Thermusthermophilus respectively.
These generally do not have 3′-5′ proof-reading ac­tivity. Polymerases are also available from other genera of bacteria (including archaebacteria), and many of these enzymes have 3′-5′ proof-reading activity (which also means, they do not usually add terminal nucleotides that are not template directed).
Proof-reading en­zymes include Tli from Thermococcuslitoraiis and Pfu from Pyrococcusfuriosus. These marine bacteria generally grow at even higher temperatures than Thermusaquaticus, and the polymerases are more ther­mo-stable than the Taq enzyme.
(d) Deoxy Nucleotide Triphosphates:
A supply of four deoxynucleotide triphos­phates, dATP, dCTP, dGTP and dTTP, are needed by the polymerase to make the new DNA.
(e) PCR Machine:
Finally we need a PCR machine to keep changing the tempera­ture. The PCR process requires cycling through sev­eral different temperatures. Because of this, PCR machines are sometimes called thermo-cyclers.
4. The PCR Reaction Cycle:
The PCR is a chain reaction because newly synthesized DNA strands will acts as template for further DNA synthesis in subsequent cycle. After 25 cycles of DNA synthesis, the products of the PCR will include, in addition to the start­ing DNA, about 10 5 copies of the specific tar­get sequence.
PCR consists of a series of cycles of three successive reactions:
This step is the first regular cycling event and consists of heating the reaction to 94-98°C for 20-30 seconds. It causes the melting of the DNA template by disrupting the hydrogen bonds between complementary bases, yielding single-stranded DNA molecules.
The reaction tempera­ture is lowered to 50-65°C for 20-40 seconds allowing annealing of the primers to the single- stranded DNA template. Typically the anneal­ing temperature is about 3-5 degrees Celsius below the Tm of the primers used.
The Tm can be determined experimentally or calculate from the following formula:
Tm = (4 x [G + C]) + (2 x [A + T])°C
Stable DNA-DNA hydrogen bonds are only formed when the primer sequence very closely matches the template sequence. The poly­merase binds to the primer-template hybrid and begins DNA synthesis.
The tem­perature at this step depends on the DNA poly­merase used. Taq polymerase has its opti­mum activity temperature at 75-80°C, and commonly a temperature of 72°C is used with this enzyme. At this step the DNA polymerase synthesizes a new DNA strand complementary to the DNA template strand by adding dNTPs that are complementary to the template in 5′ to 3′ direction.
The extension time depends both on the DNA polymerase used and on the length of the DNA fragment to be amplified. As a rule-of-thumb, at its optimum temperature, the DNA polymerase will polymerize a thousand bases per minute.
Under optimum conditions, i.e., if there are no limitations due to limiting substrates or reagents, at each extension step, the amount of DNA target is doubled, leading to exponential (geometric) amplification of the specific DNA fragment.
For example, if one starts with a single double-stranded DNA molecule, after 20 cycles the number of mol­ecules synthesized by PCR becomes 1吆 6 , and after 30 cycles the number of the DNA mol­ecules increases to 1吆 9 .
This number can be calculated by the help of following formula:
where Mf is the final number of DNA mol­ecules produced by PCR, Mf is the initial amount of DNA molecules, and n is the num­ber of PCR cycles.
This single step is oc­casionally performed at a temperature of 70- 74 °C for 5-15 minutes after the last PCR cycle to ensure that any remaining single-stranded DNA is fully extended.
5. Analysis of PCR Products:
PCR is often used as a technique to gain infor­mation about the DNA template carrying a specific target sequence. Biotechnology re­search widely depends upon PCR at various situations. Hence there are several methods for analysing the products of PCR.
Following three techniques are important:
(a) Gel Electrophoresis of PCR Products:
The final results of most PCR experiments are confirmed by subjecting a portion of the amplified reaction mixture to agarose gel electrophoresis. This can inform us the validity of the PCR experiment. If the ex­pected band during gel visualization is absent, or if additional bands are present, something has gone wrong and the experi­ment must be repeated.
In some cases, agarose gel electrophoresis is used not only to determine whether a PCR experiment has worked, but also to obtain additional information. We can also determine the presence of restriction site in the template DNA by subjecting the PCR product to re­striction endonuclease prior to electro­phoresis.
This protocol is a type of restric­tion fragment length polymorphism (RFLP) analysis which has immense significance in the construction of genome maps and studying genetic diseases. Electrophoretic analysis of PCR product can also help us in identifying the insertion or de­letion mutation in the amplified region.
(b) Cloning of PCR Products:
During the cloning experiment many times we take the help of PCR directly. If the gene of in­terest is in a very less quantity, then we need to amplify it. This is done by the help of PCR which produces enough copies of our target DNA so that we can afford to start the experiment.
(c) Sequencing of PCR Products:
The se­quencing of PCR product is done by the help of an automated sequencer machine. There is no need for radio-la­belling and autoradiography. This is an im­proved way to sequence DNA because of its speed and because it can be analysed by computer rather than a person.
PCR has a number of applications especially where speed and the number of samples to be processed are important or where the amount of DNA available is very limited. Here are some of the applications.
A. DNA Sequencing:
PCR in the presence of di-deoxynucleoside triphosphates (ddNTPs), used for DNA sequencing, al­lows DNA sequencing reactions to be run successfully with very small amounts of template.
PCR is useful as a diagnostic tool, e.g., in the identification of specific genetic traits or for the detection of patho­gens or food contaminants. One of the first applications of PCR to genetic diagnosis was for sickle cell anaemia.
The ability to amplify DNA from regions of the genome that are highly polymorphic (and which are variable be­tween individuals) starting with samples containing very small amounts of DNA (e.g., single hairs or traces of body fluids, such as blood and semen) leads to applica­tions in forensic work.
D. Present-Day Population Genetics:
It al­lows for the determination of frequencies of particular alleles in a large collection of individuals. A particular advantage of us­ing PCR in population genetic studies is that, with appropriately designed specific primers, it may be possible to amplify DNA from one organism that cannot be sepa­rated from others, such as a particular bacterial strain in a mixed population.
(Such primers will anneal to the target DNA from the organism of interest, but not to DNA from others.)
E. Archaeology and Evolution:
PCR can be used with old material as well as more re­cent samples, and it is often possible to amplify ancient DNA from museum speci­mens and archaeological remains. Mostly mitochondrial DNA or chloroplast DNA is used. This allows inferences to be made about the origins of particular populations or species.
7. Precautions and Drawbacks:
The size of fragments that can be am­plified is limited by the processivity of the polymerase used. Using a mix­ture of polymerases that includes a proof­reading enzyme increases the size of pro­duct that can be obtained (up to 10 kbp or more), because incorrectly incorporated nucleotides can be removed rather than causing chain termination.
Ii. Amplifying the Wrong Sequence:
PCR depends on the ability of the primers to anneal to the correct sequence, and this depends on the conditions of annealing (ionic concentration, temperature, etc.) and the actual sequence (or sequences if mixed sites are included) of the primers.
It is possible for primers to anneal to the “wrong” part of the target DNA, through chance complementarity. If this happens and the primers anneal in the correct ori­entation to each other (i.e., directing syn­thesis towards each other) and at sites that are not too far apart, then the result is the amplification of a sequence other than the desired one.
The possibility of incorrect annealing may be avoided by use of longer primers, which will be more specific in their annealing sites. Raising the temperature and adjust­ing the concentration of magnesium ions (which stabilize primer-template binding) can be used to increase the specificity of primer binding.
Because of the extraor­dinary sensitivity of PCR, there is a par­ticular danger of contaminating the DNA sample to be amplified with extraneous material. This is particularly important when using material containing only small amounts of DNA, as with archaeological work.
Contamination might be of labora­tory origin (e.g., from aerosols created by pipetting solutions containing related DNA sequences, including material amplified previously by PCR) or of external ori­gin (perhaps by bacterial, fungal or human contamination of sample tissue).
Laboratory contamination can be mini­mized by precautions such as careful use and design of pipettes, separation of the pre-PCR and post-PCR stages of an experi­ment into different rooms. Contamination from other sources can be reduced by care­ful handling and preparation of a sample before amplification.
Iv. Sequence Heterogeneity:
Amplification may give rise to a mixture of molecules of slightly different sequences.
A mixture could arise for several reasons:
If the template DNA came from an individual heterozygous at the locus in question, each of the alleles present should be represented in similar quantities in the PCR prod­ucts.
(b) Population Heterogeneity:
If the tem­plate DNA came from several individu­als rather than a single one, heteroge­neity in the population may give rise to heterogeneity in the products.
(c) DNA Damage and Polymerase Error:
Heterogeneity can also arise from damage to DNA before amplification, especially if the sample has not been carefully preserved. Therefore, this is particularly likely to be a problem with archaeological and forensic material.
V. Jumping PCR:
When degraded DNA is amplified, it may be that any given sample molecule is not long enough to span the entire distance between the two priming sites. The result in the first round of syn­thesis would be extension of the primer to the end of a fragmented molecule, but not all the way to the second primer site.
How­ever, on a subsequent round of synthesis, the truncated amplification product may anneal to a different DNA fragment that contains the remaining region intact. This would then allow synthesis of the full PCR product. This is called jumping PCR. So it is sometimes possible to generate PCR products that are longer than any indi­vidual template molecule. This can be ad­vantageous when amplifying badly de­graded DNA.
A. Hot-Start PCR:
As soon as the PCR re­agents have all been mixed together, it is possible for the DNA polymerase to start synthesis. This may happen while the re­action mixture is being heated for the first time, and is at a temperature low enough to allow non-specific annealing of primer to template, generating a range of non-spe­cific products.
This problem would be pre­vented if DNA synthesis could not take place until the first cycle had reached its maximum temperature. This is the basis of hot-start PCR. In the simplest form, the DNA polymerase is not added to the reaction tubes until they have reached the DNA melting temperature of the first cycle. This is satisfactory where small numbers of samples are being processed, but not with large numbers.
B. Touch-Down PCR:
The annealing tem­perature used in conventional PCR is usu­ally several degrees below the maximum at which primers can remain bound to tem­plate, to ensure stable binding. However, this use of a lower temperature permits a small amount of mismatching between primers and template, which may allow primers to bind to incorrect sites and gen­erate spurious products.
The effects of this can be reduced with touch-down PCR. In this, a high annealing temperature is used initially (at which even correct bind­ing may not be possible). The annealing temperature is reduced in subsequent rounds. There will, therefore, come a point at which correctly matched primer-tem- plate annealing is just possible, but incor­rect matching is not and the desired prod­ucts will be the most abundant.
C. Nested PCR:
Here, two successive PCRs are carried out. The first PCR uses a con­ventional template. The products of the first PCR are then used as the template for the second PCR, with primers that are designed to anneal within the desired prod­uct of the first PCR.
Although the first PCR may generate some non-specific products in addition to the desired products, it is unlikely that the non-specific products will also contain annealing sites for both the primers used in the second PCR. Thus, only the desired products from the first PCR are likely to be suitable templates for the second.
D. Inverse PCR:
It is possible to arrange for the amplification of sequences outside the primers, in a technique called inverse PCR (IPCR). In this technique the sample DNA is first cut with an enzyme outside the region whose sequence is already known.
The resulting linear molecules are then circularized, by ligation under condi­tions that favour intermolecular reactions. A second restriction digestion is then done, using an enzyme cutting within the region of known sequence.
The result is now that the first fragment containing this sequence has been turned ‘inside out’, leaving known sequence on the outside and the material that had previously been flanking it within. Primers complementary to the known sequence on the outside of the mole­cule can now be used to amplify the region of interest between them.
E. Reverse Transcriptase PCR:
It is often convenient to amplify RNA molecules, per­haps as a precursor to cloning them, or to estimate the abundance of a particular mRNA in a sample. This is usually done by having a round of reverse transcription, using a reverse transcriptase enzyme and a single primer, to make a single strand of cDNA prior to the PCR itself.
The primer for reverse transcription could be oligo-dT for general cDNA synthesis from polyadenylated messages, or it could be specific to a particular message.
F. In Situ PCR:
It is possible to carry out PCR using permeabilized tissue, such as thin sections on a microscope slide. This requires a specially adapted PCR machine to accommodate the slide. If the PCR prod­uct can be detected (perhaps by hybridiza­tion, also in situ), then this allows one to identify where in the tissue the target nucleic acid is located.
G. Asymmetric PCR:
By reducing the amount of one of the two primers, it is pos­sible to arrange for preferential amplifica­tion of one of the strands, resulting in a preparation of single-stranded DNA, which has a number of uses in molecular biology. Preferential amplification of one strand in this way is known as asymmet­ric PCR.
H. Anchored PCR:
Anchored PCR is applied when only one piece of sequence (and therefore, one priming site) for the region of interest is known. The aim is to attach the region to be amplified to a piece of known sequence and then to use that as the second priming site.
There are two ways in which this can most easily be done. One is to fragment the sample DNA and ligate it to molecules of known sequence, such as a vector. This known sequence is used as the basis for designing one of the two PCR primers. The second method is to add tails enzymatically to the sample DNA or the molecules produced after the first round of synthesis.
I. Emulsion PCR:
In a conventional PCR, the reactions are carried out inside plastic tubes. It is possible to incorporate all the reagents inside lipid droplets and carry out PCR on a much smaller scale. This has certain advantages. It is possible to in­crease and decrease the temperature of small droplets very quickly.
In addition, if each droplet contains a single template molecule at the start, then all the products in an individual droplet result from the am­plification of a single template molecule. The method is also called droplet PCR.
J. Isothermal Amplification:
The repeated heating and cooling required by PCR lim­its how quickly the process can be carried out. Loop-mediated isothermal ampli­fication (LAMP) has been developed, which allows templates to be amplified at a constant temperature (typically around 65°C).
It uses a DNA polymerase with strand-displacing activity and avoids the need for heating to high temperatures. This method is used for the detection of pathogens outside of specialist laboratories.
K. Real Time PCR:
It is possible to use PCR to estimate the abundance of a particular nucleic acid molecule in a sample. This can be done by real time PCR. This can be done in two ways.
In the first, a fluorescent, double-stranded DNA (dsDNA)-binding dye (such as SYBR green) is present in the PCR. As dsDNA prod­uct accumulates, the amount of fluorescence from the dye increases, and this can be de­tected.
The experiment requires a PCR ma­chine that is also equipped with a fluorescence measurement facility. Because the method simply detects dsDNA, it measures the amount of PCR product at a given time regardless of whether it is from the correct region.
The second approach to real-time PCR al­lows detection of a specific product, rather than dsDNA in general, and uses a specially syn­thesized probe oligonucleotide. This probe is designed to anneal within the region to be amplified and carries a fluorescent reporter dye at one end and a quencher at the other end of the molecule.
If the quencher and the reporter are in close proximity (i.e., attached to the same oligonucleotide), then the quencher stops the reporter from fluorescing.
During PCR, the probe will anneal to single-stranded DNA within the target region. When the poly­merase meets the annealed probe, the 5′-3′ exonuclease activity of the enzyme degrades the probe, liberating the reporter from the quencher. Thus, the fluorescent reporter ac­cumulates during the course of the PCR. This type of PCR mechanism is shown in the dia­gram given above.
Question : PCR Below, you see: Figure A: DNA sequence (800 nt from the first ATG to the last TAA) with positions of restriction enzymes’ sites Figure B: the multiple cloning sites (MCS) of the plasmid DNA, and Figure C: the activities of the listed restriction enzymes in three common buffers. Task: You will need to find a strategy to introduce the coding region of the
Figure C: the activities of the listed restriction enzymes in three common buffers.
Task: You will need to find a strategy to introduce the coding region of the gene (from the first ATG to the last TAA) into the multiple cloning sites (MCS) of the vector using PCR and molecular cloning techniques.
For this task you will need:
(Task 1) to select two restriction enzymes in MCS to introduce your DNA (5 marks)
(Task 2) to design forward and reverse PCR primers, which contain the sequences of selected restriction sites (10 marks for each primer = 20 marks total)
(Task 3) Calculate Tm of both primers (5 marks for each, 10 marks total).
Site Directed Mutagenesis
Site-directed mutagenesis (SDM) is a method to create specific, targeted changes in double stranded plasmid DNA. There are many reasons to make specific DNA alterations (insertions, deletions and substitutions), including:
- To study changes in protein activity that occur as a result of the DNA manipulation.
- To select or screen for mutations (at the DNA, RNA or protein level) that have a desired property
- To introduce or remove restriction endonuclease sites or tags
SDM is an in vitro procedure that uses custom designed oligonucleotide primers to confer a desired mutation in a double-stranded DNA plasmid. Formerly, a method pioneered by Kunkel (Kunkel, 1985) that takes advantage of a strain deficient in dUTPase and uracil deglycosylase so that the recipient E. coli degrades the uracil-containing wild-type DNA was widely used. Currently, there are a number of commercially available kits that also require specific modification and/or unique E. coli strains (for example, the Phusion Site-Directed Mutagenesis ® from Thermo and the GeneArt ® system from Life). The most widely-used methods do not require any modifications or unique strains and incorporate mutations into the plasmid by inverse PCR with standard primers. For these methods, primers can be designed in either an overlapping (QuikChange ® , Agilent) or a back-to-back orientation (Q5 ® Site-Directed Mutagenesis Kit) (Figure 1). Overlapping primer design results in a product that will re-circularize to form a doubly-nicked plasmid. Despite the presence of these nicks, this circular product can be directly transformed into E. coli, albeit at a lower efficiency than non-nicked plasmids. Back-to-back primer design methods not only have the advantage of transforming non-nicked plasmids, but also allow exponential amplification to generate significantly more of the desired product (Figure 2). In addition, because the primers do not overlap each other, deletions sizes are only limited by the plasmid and insertions are only limited by the constraints of modern primer synthesis. Currently, by splitting the insertion between the two primers, insertions up to 100 bp can routinely be created in one step using this method.
Before primers are designed, it is important to determine which mutagenesis workflow is to be used. Here we present a comparison of three commercially available kits (Figure 3) and a brief description of important features.
Before you plan your next SDM experiment, be sure to read through our list of important experimental considerations.
Figure 1: Site-specific mutagenesis proceeds in less than 2 hours
The use of a master mix, a unique multi-enzyme KLD enzyme mix, and a fast polymerase ensures that, for most plasmids, the mutagenesis reaction is complete in less than two hours.
Figure 2: Q5 Site-Directed Mutagenesis Kit Overview
This kit is designed for rapid and efficient incorporation of insertions, deletions and substitutions into doublestranded plasmid DNA. The first step is an exponential amplification using standard primers and a master mix fomulation of Q5 Hot Start High-Fidelity DNA Polymerase. The second step involves incubation with a unique enzyme mix containing a kinase, a ligase and DpnI. Together, these enzymes allow for rapid circularization of the PCR product and removal of the template DNA. The last step is a high-efficiency transformation into chemicallycompetent cells (provided).
Figure 3: Primer Design for the Q5 Site-Directed Mutagenesis Kit
Cloning-free template DNA preparation for cell-free protein synthesis via two-step PCR using versatile primer designs with short 3'-UTR
Cell-free protein synthesis (CFPS) systems largely retain the endogenous translation machinery of the host organism, making them highly applicable for proteomics analysis of diverse biological processes. However, laborious and time-consuming cloning procedures hinder progress with CFPS systems. Herein, we report the development of a rapid and efficient two-step polymerase chain reaction (PCR) method to prepare linear DNA templates for a wheat germ CFPS system. We developed a novel, effective short 3'-untranslated region (3'-UTR) sequence that facilitates translation. Application of the short 3'-UTR to two-step PCR enabled the generation of various transcription templates from the same plasmid, including fusion proteins with N- or C-terminal tags, and truncated proteins. Our method supports the cloning-free expression of target proteins using an mRNA pool from biological material. The established system is a highly versatile platform for in vitro protein synthesis using wheat germ CFPS.
Keywords: 3′-UTR cell-free protein synthesis wheat germ extract.
© 2017 Molecular Biology Society of Japan and John Wiley & Sons Australia, Ltd.
Colony PCR is a convenient high-throughput method for determining the presence or absence of insert DNA in plasmid constructs. Individual transformants can either be lysed in water with a short heating step or added directly to the PCR reaction and lysed during the initial heating step. This initial heating step causes the release of the plasmid DNA from the cell, so it can serve as template for the amplification reaction. Primers designed to specifically target the insert DNA can be used to determine if the construct contains the DNA fragment of interest. Alternatively, primers targeting vector DNA flanking the insert can be used to determine whether or not the insert is the correct molecular size. Insert specific primers can provide information on both the specificity and size of the insert DNA while the use of vector specific primers allows screening of multiple constructs simultaneously. Colony PCR can also be used to determine insert orientation. PCR amplification of the plasmid using an insert specific primer paired with a vector specific primer can be designed to produce an amplicon of a specific size only if the insert is in the correct orientation. In all experimental designs, presence or absence of a PCR amplicon and size of the product are determined by electrophoresis alongside a DNA size marker on an agarose gel.
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The gene of interest usually has to be amplified from genomic or vector DNA by PCR (polymerase chain reaction) before it can be cloned into an expression vector. The first step is the design of the necessary primers.
Primer sequence. Especially the 3'-end of the primer molecule is critical for the specificity and sensitivity of PCR. It is recommended not to have:
- 3 of more G or C bases at this position. This may stabilize nonspecific annealing of the primer.
- a 3' thymidine, since it is more prone to mispriming than the other nucleotides.
Primer pairs should be checked for complementarity at the 3'-end. This often leads to primer-dimer formation.
Bases at the 5'-end of the primer are less critical for primer annealing. Therefore, it is possible to add sequence elements, like restriction sites, to the 5'-end of the primer molecule.
Primer length. Usually a primer length of 18-30 bases is optimal for most PCR applications. Shorter primers could lead to amplification of nonspecific PCR products.
Melting temperature (Tm). The specificity of PCR depends strongly on the melting temperature (Tm) of the primers (the temperature at which half of the primer has annealed to the template). Usually good results are obtained when the Tm's for both primers are similar (within 2-4 °C) and above 60°C. The Tm for a primer can be estimated using the following formula:
GC content. The GC content of a primer should be between 40 and 60%.
Design of the 5'-end primer
The 5'-end primer overlaps with the 5'-end of the gene of interest and should contain the following elements:
- Restriction site. The restriction site should be the same or provide the same sticky end to the first of the restriction enzymes in the multiple cloning site of the vector chosen to clone the gene of interest into. Alternatively, you could pick any restriction enzyme that gives a blunt end upon cleavage (see cloning). Often Nco ICCATGG) or Nde I (CATATG) are chosen because the ATG within these sites can be used directly to create the ATG start codon and/or the ATG codon for the N-terminal methionine residue (see Utilisation of the Nco I cloning site)
- 5'-extension to the restriction site. Restriction enzymes cleave DNA much less efficient towards the end of a fragment. A 5' extension of the restriction site with 2-10 bases greatly increases the cleavage efficiency of most enzymes. Data on the effect of the extension length and sequence on the cleavage efficiencies of the most used restriction enzymes can be found in the reference appendix of the New England Biolabs catalogue.
- Start codon. A start codon (usually ATG) should be included when the gene of interest is not expressed with an N-terminal tag or fusion partner or when an N-terminal methionine residue is present. It should be checked that the start codon and the gene of interest are in frame with an eventual N-terminal tag and/or fusion partner.
- Overlap with the gene of interest. The overlap between the primer and the gene of interest should be long enough to give a Tm of 60°C or more (calculated as shown above).
8 Approaches to Random Mutagenesis
Random mutagenesis is an incredibly powerful tool for altering the properties of enzymes. Imagine, for example, you were studying a G-protein coupled receptor (GPCR) and wanted to create a temperature-sensitive version of the receptor or one that was activated by a different ligand than the wild-type. How could you do this?
Firstly, you would clone the gene encoding the receptor, then randomly introduce mutations into the gene sequence to create a “library” containing thousands of versions of the gene. Each version (or “variant”) of the gene in the library would contain different mutations and so encode receptors with slightly altered amino acid sequences giving them slightly different enzymatic properties than the wild-type.
Next, you could transform the library into a strain where the receptor would be expressed and apply a high throughput screen to pick out variants in the library that have the properties you are looking for. Using a high throughput screen for GPCR activity (see here for examples) you could pick out the variants from the library that were temperature-sensitive or were activated by different ligands.
Sound easy? Well, of course it’s not that easy. Creating a random mutant library that contains enough variants to give you a good chance of obtaining the altered enzyme you desire is a challenge in itself. There are many ways to create random mutant libraries, each with it’s own pros and cons. Here are some of them:
1. Error-prone PCR. This approach uses a “sloppy” version of PCR, in which the polymerase has a fairly high error rate (up to 2%), to amplify the wild-type sequence. The PCR can be made error-prone in various ways including increasing the MgCl2 in the reaction, adding MnCl2 or using unequal concentrations of each nucleotide. Here is a good review of error prone PCR techiques and theory. After amplification, the library of mutant coding sequences must be cloned into a suitable plasmid. The drawback of this approach is that size of the library is limited by the efficiency of the cloning step. Although point mutations are the most common types of mutation in error prone PCR, deletions and frameshift mutations are also possible. There are a number of commercial error-prone PCR kits available, including those from Stratagene and Clontech.
2. Rolling circle error-prone PCR is a variant of error-prone PCR in which wild-type sequence is first cloned into a plasmid, then the whole plasmid is amplified under error-prone conditions. This eliminates the ligation step that limits library size in conventional error-prone PCR but of course the amplification of the whole plasmid is less efficient than amplifying the coding sequence alone. More details can be found here.
3. Mutator strains. In this approach the wild-type sequence is cloned into a plasmid and transformed into a mutator strain, such as Stratagene’s XL1-Red. XL1-red is an E.coli strain whose deficiency in three of the primary DNA repair pathways (mutS, mutD and mutT) causes it to make errors during replicate of it’s DNA, including the cloned plasmid. As a result each copy of the plasmid replicated in this strain has the potential to be different from the wild-type. One advantage of mutator strains is that a wide variety of mutations can be incorporated including substitutions, deletions and frame-shifts. The drawback with this method is that the strain becomes progressively sick as it accumulates more and more mutations in it’s own genome so several steps of growth, plasmid isolation, transformation and re-growth are normally required to obtain a meaningful library.
4. Temporary mutator strains. Temporary mutator strains can be built by over-expressing a mutator allele such as mutD5 (a dominant negative version of mutD) which limits the cell’s ability to repair DNA lesions. By expressing mutD5 from an inducible promoter it is possible to allow the cells to cycle between mutagenic (mutD5 expression on) and normal (mutD5 expression off) periods of growth. The periods of normal growth allow the cells to recover from the mutagenesis, which allows these strains to grow for longer than conventional mutator strains.
If a plasmid with a temperature-sensitive origin of replication is used, the mutagenic plasmid can easily be removed restore normal DNA repair, allowing the mutants to be grown up for analysis/screening. An example of the construction and use of such a strain can be found here. As far as I am aware there are no commercially available temporary mutator strains.
5. Insertion mutagenesis. Finnzymes have a kit that uses a transposon-based system to randomly insert a 15-base pair sequence throughout a sequence of interest, be it an isolated insert or plasmid. This inserts 5 codons into the sequence, allowing any gene with an insertion to be expressed (i.e. no frame-shifts or stop codons are cause). Since the insertion is random, each copy of the sequence will have different insertions, thus creating a library.
6. Ethyl methanesulfonate (EMS) is a chemical mutagen. EMS aklylates guanidine residues, causing them to be incorrectly copied during DNA replication. Since EMS directly chemically modifies DNA, EMS mutagenesis can be carried out either in vivo (i.e. whole-cell mutagenesis) or in vitro. An example of in vitro mutagenesis with EMS in which a PCR-amplified gene was subjected to reaction with EMS before being ligated into a plasmid and transformed can be found here.
7. Nitrous acid is another chemical mutagen. It acts by de-aminating adenine and cytosine residues (although other mechanisms are discussed here) causing transversion point mutations (A/T to G/C and vice versa). An example of a study using nitrosoguanidine mutagenesis can be found here.
Note: I have only mentioned two chemical mutagens but there are many others. Hirokazu Inoue has written an excellent article describing some of them and their use in mutagenesis, see here (pdf).
Another note: Chemical mutagens are, of course… mutagens and therefore should be handled with great care. Be especially careful with EMS as it is volatile at room temperature. Read the MSDS and do a proper risk assessment before carrying out these experiments.
What are the Similarities Between Forward and Reverse Primer?
- Both Forward and Reverse primers are made from oligonucleotides.
- Both Forward and Reverse Primers possess short nucleotide sequence complementary to the flanking ends of the DNA double strands.
- Both Forward and Reverse primers usually consist of 20 nucleotides.
- Both Forward and Reverse Primers are used in polymerase chain reactions.
- Both Forward and Reverse Primers are synthesized commercially.
- Both Forward and Reverse primers are temperature stable and they are normally having similar Tm.
- Both Forward and Reverse Primers are annealed with the target DNA sequences.
- Both Forward and Reverse Primers are designed according to the PCR reactions.
- Both Forward and Reverse Primers are served as starting points for the DNA amplification.
- Both reverse and forward primers are important for the production of million copies of particular regions of targeted or interested DNA sequences.
Uses of PCR
PCR can be used for a broad variety of experiments and analyzes. Some examples are discussed below.
Genetic fingerprinting is a forensic technique used to identify a person by comparing his or her DNA with a given sample, e.g., blood from a crime scene can be genetically compared to blood from a suspect. The sample may contain only a tiny amount of DNA, obtained from a source such as blood, semen, saliva, hair, etc. Theoretically, just a single strand is needed. First, one breaks the DNA sample into fragments, then amplifies them using PCR. The amplified fragments are then separated using gel electrophoresis. The overall layout of the DNA fragments is called a DNA fingerprint.
Although these resulting 'fingerprints' are unique (except for identical twins), genetic relationships, for example, parent-child or siblings, can be determined from two or more genetic fingerprints, which can be used for paternity tests (Fig. 4). A variation of this technique can also be used to determine evolutionary relationships between organisms.
Detection of hereditary diseases
The detection of hereditary diseases in a given genome is a long and difficult process, which can be shortened significantly by using PCR. Each gene in question can easily be amplified through PCR by using the appropriate primers and then sequenced to detect mutations.
Viral diseases, too, can be detected using PCR through amplification of the viral DNA. This analysis is possible right after infection, which can be from several days to several months before actual symptoms occur. Such early diagnoses give physicians a significant lead in treatment.
Cloning a gene--not to be confused with cloning a whole organism--describes the process of isolating a gene from one organism and then inserting it into another organism (now termed a genetically modified organism (GMO)). PCR is often used to amplify the gene, which can then be inserted into a vector (a vector is a piece of DNA which 'carries' the gene into the GEO) such as a plasmid (a circular DNA molecule) (Fig. 5). The DNA can then be transferred into an organism (the GMO) where the gene and its product can be studied more closely. Expressing a cloned gene (when a gene is expressed the gene product (usually protein or RNA) is produced by the GMO) can also be a way of mass-producing useful proteins--for example, medicines or enzymes in biological washing powders. The incorporation of an affinity tag on a recombinant protein will generate a fusion protein which can be more easily purified by affinity chromatography.
Mutagenesis is a way of making changes to the sequence of nucleotides in the DNA. There are situations in which one is interested in mutated (changed) copies of a given DNA strand, for example, when trying to assess the function of a gene or in in-vitro protein evolution. Mutations can be introduced into copied DNA sequences in two fundamentally different ways in the PCR process. Site-directed mutagenesis allows the experimenter to introduce a mutation at a specific location on the DNA strand. Usually, the desired mutation is incorporated in the primers used for the PCR program. Random mutagenesis, on the other hand, is based on the use of error-prone polymerases in the PCR process. In the case of random mutagenesis, the location and nature of the mutations cannot be controlled. One application of random mutagenesis is to analyze structure-function relationships of a protein. By randomly altering a DNA sequence, one can compare the resulting protein with the original and determine the function of each part of the protein.
Analysis of ancient DNA
Using PCR, it becomes possible to analyze DNA that is thousands of years old. PCR techniques have been successfully used on animals, such as a forty-thousand-year-old mammoth, and also on human DNA, in applications ranging from the analysis of Egyptian mummies to the identification of a Russian tsar.
Genotyping of specific mutations
Through the use of allele-specific PCR, one can easily determine which allele of a mutation or polymorphism an individual has. Here, one of the two primers is common, and would anneal a short distance away from the mutation, while the other anneals right on the variation. The 3' end of the allele-specific primer is modified, to only anneal if it matches one of the alleles. If the mutation of interest is a T or C single nucleotide polymorphism (T/C SNP), one would use two reactions, one containing a primer ending in T, and the other ending in C. The common primer would be the same. Following PCR, these two sets of reactions would be run out on an agarose gel, and the band pattern will tell you if the individual is homozygous T, homozygous C, or heterzygous T/C. This methodology has several applications, such as amplifying certain haplotypes (when certain alleles at 2 or more SNPs occur together on the same chromosome [Linkage Disequilibrium]) or detection of recombinant chromosomes and the study of meiotic recombination.
Comparison of gene expression
Researchers have used traditional PCR as a way to estimate changes in the amount of a gene's expression. Ribonucleic acid (RNA) is the molecule into which DNA is transcribed prior to making a protein, and those strands of RNA that hold the instructions for protein sequence are known as messenger RNA (mRNA). Once RNA is isolated it can be reverse transcribed back into DNA (complementary DNA to be precise, known as cDNA), at which point traditional PCR can be applied to amplify the gene, this methodology is called RT-PCR. In most cases if there is more starting material (mRNA) of a gene then during PCR more copies of the gene will be generated. When the product of the PCR reaction are run on an agarose gel (see Figure 3 above) a band, corresponding to a gene, will appear larger on the gel (note that the band remains in the same location relative to the ladder, it will just appear fatter or brighter). By running samples of amplified cDNA from differently treated organisms one can get a general idea of which sample expressed more of the gene of interest. A quantative RT-PCR method has been developed, it is called Real-Time PCR.