UNIT 8: POLYMERASE CHAIN REACTION (PCR)
INTRODUCTION:
PCR has completely revolutionized how most DNA cloning experiments are done. Whether you are trying to identify a gene, clone it, express it, or mutagenize it, there is probably a PCR method available. PCR is the only common technique with its own journal-cleverly named PCR. Two somewhat out-of-date texts which discuss a wide variety of PCR techniques are:
PCR Protocols: A Guide to Methods and Applications, Academic Press, (1990)
PCR Technology Principles and Applications for DNA Amplification , Stockton Press, (1989)
A more recent book is The Polymerase Chain Reaction, K. Mullis et al. Birkhäuser, Boston (1994)
Ausubel et al. sections 15.0.3 - 15.7.6 also contains useful information.
To understand the PCR reaction you must first know the following information:
1. The strands of a DNA double helix run in an opposite polarity along the backbone such that if one strand is viewed as extending in the 5' to 3' direction, the other strand runs in the 3' to 5' direction.
2. DNA dependent DNA polymerases polymerize dNTPs in presence of a divalent cation onto the 3' end of the growing DNA chain. Note that the enzyme requires a primer DNA or RNA terminated by a 3'-OH group hybridized to the template DNA in order to initiate synthesis on a DNA template. See diagram below.
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3. An oligonucleotide of about 15-17 bases will normally hybridize to a unique site on a DNA even as large as the human genome (3 x 109 bp).
A PCR reaction consists of three steps which are repeated multiple times: denaturation, annealing, and synthesis. These steps are incorporated into a PCR reaction as illustrated in the diagram below.
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The PCR reaction mix consists of:
1. The template DNA
2. Two primers which are complementary to the opposite strands and which flank the region to be amplified
3. dNTPs (the standard deoxyribonucleoside triphosphates, i.e. dATP, dCTP, dGTP, and dTTP)
4. Buffer and salts
5. Thermostable polymerase
In the first step of a PCR reaction the DNA is denatured by heating to 93oC to 94oC so that the strands come apart. In the second step, the reaction is cooled to a lower temperature, typically 50oC to 72oC, such that the primers anneal selectively to the regions which flank the target site to be amplified. Lastly, the reaction is heated to 72oC to 75oC and the DNA polymerase extends the primers such that a copy of the target region is synthesized. Typically this three step sequence is repeated for 25 to 35 cycles. Additional cycles, however, will often generate nonspecific products.
Commonly, oligonucleotides of about 20 nucleotides long are used as primers. Primers should not contain a sequence with any internal secondary structures, especially at the 3' end because the oligonucleotide may not anneal properly. In addition the primers should not be able to hybridize to each other in order to avoid the synthesis of primer dimers. Runs of more than three bases of a single nucleotide are also best avoided because they can anneal to repeated regions in the genome. With any primer it is desirable to confirm that the sequence is not found in other parts of the genome. This can be done by comparing the sequence to the sequences available in Genbank database. The G+C content of the primer will affect the reaction temperature. It is therefore desirable to make the G+C contents of the two primers similar. Computer programs are available that assist in the process of primer design.
There are a number of commercially available thermostable DNA polymerases. Taq polymerase (Perkin Elmer, Roche Biomolecular - formally Boehringer Mannheim) is the most widely used. It has a moderate error rate and is good for amplifying sequences of 2000 bp or less. KlenTaq LA (Ab Peptides) is highly processive and able to amplify targets up to 18 kb. The error rate is significantly lower than native Taq. Hot Tub (Amersham) is highly processive and has been used to amplify sequences up to 18 kb. It's error rate is slightly less than Taq. Pfu (Stratagene) is not very processive and is only good for amplifying short sequences, typically less than 1 kb. However it has the lowest error rate reported to date. Turbo Pfu is an improved version of Pfu that is capable of amplifying relatively long regions (~10Kb) with high fidelity.
One of the most exciting new developments in PCR technology is Long PCR (W. Barnes, Proc. Natl. Acad. Sci. 91: 2216-2220 (1994). By using a combination of thermostable polymerases, Barnes has recently been able to achieve amplifications of as much as 50 kb of template.
The reaction condition for each experiment may vary and has to be optimized to obtain the best results. The reaction temperature is the easiest to change. The annealing temperature must be low enough to get sufficient annealing to start the PCR reaction efficiently, but high enough to prevent nonspecific priming. Some primer sets also require altering the Mg++ concentration or addition of DMSO (up to 15 %) or Formamide (up to 10%) to increase specificity. Note in particular that the optimum reaction conditions often differ for different DNA polymerases, different types of tubes e.g. regular vs. thin wall, and different PCR machines. Unfortunately, the optimum conditions have to be determined empirically.
“Hot-Start”
A commonly used procedure is the "hot start". A hot start is carried out by adding wax to an incomplete PCR reaction mix lacking at least one component needed for the reaction e.g. the polymerase, then going through one cycle and finally cooling the tubes down. This procedure melts the wax, allows it to float to the top, and then allows it to resolidify at the top of the reaction mix. The missing component e.g. polymerase is then placed on top of the solidified wax where it will be released when the samples reach a temperature near 70oC. The release of the missing component only after the sample is hot prevents synthesis of DNA at low temperatures where illegitimate priming is more likely to occur. As a result, hot start often enhances specificity and yield. Recently several suppliers of Taq polymerase have developed versions of Taq in which the enzyme is supplied as an inactive complex with an anti-Taq antibody. The enzyme becomes active only when the antibody is denatured by heating to 90oC or above for several minutes. After the enzyme is heat activated it begins to work when primers anneal as the temperature drops at the start of the first cycle. Because the primers that bind at the higher temperature are more likely to be bound specifically, this reduces the chances of illegitimate priming.
Sensitivity And Contamination
The beauty of PCR is it is so powerful that one can amplify from a single DNA molecule. This great sensitivity does have one great disadvantage in that one must be constantly aware of the risk of contamination. Aliquoting reagents, inclusion of multiple negative controls in the experiments, physical separation of places for setting up the reaction from those used for analyzing the results, as well as developing meticulous laboratory techniques, are some of the suggestions to reduce the risk of contamination. One of the things that helps most is to use commercially plugged tips that prevent the creation of aerosols during pipetting. In addition, it is probably a good idea to always add the DNA template as the last addition to the PCR mix. This minimizes the possibility of transferring template molecules from one tube to another when a tube must be opened and closed multiple times to assemble the reaction mix.
Typically concentrations for PCR reactions are (assuming a 50 ml reaction volume):
Template DNA (assuming genomic mammalian DNA) - 50 ng to 500 ng. If you are trying to amplify a template from an organism of less genomic complexity, you can use even less. It is not easy to give you clear guidelines about the required purity of the template DNA. Some primer pairs are able to work satisfactorily even with relatively crude template preparations. In other cases, the template may have to be extensively purified before use. We provide several protocols in this manual for the preparation of template DNAs.
NOTE: Too much template can result in the amplification of nonspecific products or even cause the complete failure of the reaction. The nonspecific reaction is due to the fact that at high template concentrations, the initial high concentration of primers can lead to nonspecific hybridization. The failure problem can be due to the fact that during the first round of synthesis the polymerase continues on down the template for a long time. The more initial templates there are, the more nucleotides are used up initially, which limits the efficiency of amplifications in the later cycles. Therefore, for each PCR reaction there is an optimal template concentration. Template concentrations too low result in poor initial priming: too high: the nucleotides triphosphates are used up.
Primers - Usually about 100-500 ng of each primer will work in most cases. Use the lowest concentration of primers that generate a good yield of your desired product. However, you may have to increase the concentration of primers if the primers are degenerate.
Just as in the case of the purity of the template DNA there is no hard and fast rule as to how pure the primers used for PCR need to be, and different workers have used many different purification methods. As usual it helps to understand what it is you are trying to remove. If you feel that the few percent of deletion oligonucleotides that typically contaminate synthetic oligonucleotide preparations are producing non-specific bands in your PCR reaction, you will need to remove them. There are two common ways of doing this. Many researchers purify primers by gel electrophoresis using 7M urea and 12% polyacrylamide. The bands on the gel can be detected by holding the gel over a fluorescent TLC plate under UV light. The deletion oligonucleotides run faster than the product band which can be cut out of the acrylamide and recovered by crushing and soaking the gel fragments overnight in water. This method can be used for primers of any size, but it is slow, requires a good deal of hands on time, and probably is not necessary in many cases. A far easier method for primers up to a length of about 25 is to use silica gel 60 thin layer (0.2mm) chromatography plates containing a fluorescent indicator (Merck) . The primers are spotted in a few ml of water and developed for about 45 minutes in a TLC chamber using as an eluting solvent n-propanol: ammonia: hydroxide:water (55:35:10). In this case the correct band is usually the darkest band closest to the origin seen using a hand-held UV (254nm) light. It is easy to scrape off the layer containing the desired band and elute the oligonucleotide in a few hundred microliters of water. After standing for a few minutes, the silica gel is spun down and the supernatant can be spin filtered to remove the last traces of silica. This method, like the gel electrophoresis method actually removes the deletion oligonucleotides as well as other impurities from the synthesis. A commercial kit based on this method is sold by Amersham.
Other workers desalt the crude primers on Sep Pack reversed phase columns (Waters) (See Protocol 8.6). While this method certainly removes organic impurities, it does nothing to remove the deletion oligonucleotides. The use of reversed phase columns makes more sense if the oligonucleotide is synthesized with the trityl protecting group left on. In this case, use of a revered phase column will remove organic impurities as well as the deletion oligonucleotides since they lack the highly hydrophobic trityl group. Subsequently the blocked oligonucleotide is eluted by decreasing the polarity of the solvent. It must then be deblocked. Because of the extra steps involved, this method is not widely used.
The simplest way of cleaning up oligonucleotides is probably precipitation and/or extraction with n-butanol. Simply take 1 part of oligonucleotide solution and mix with 9 parts of n-butanol . Vortex vigorously. Then spin the precipitate down for 15 minutes in a table top centrifuge and discard the the supernatant. (Alternatively one can add an equal volume of butanol, vortex, and remove the aqueous layer. If this is repeated two more times the solution is readily concentrated. The residual butanol can then be removed by a quick spin in a SpeedVac centrifuge .) This method can be used directly from the ammonia hydroxide deprotecting solution and is described by Sawadogo and VanDyke in Nucleic Acids Research , 19: (3) p. 674. While this method does not remove the deletion oligonucleotides, it does remove the organic impurities that might poison the PCR reaction. In our hands, this method seems quite adequate for routine use.
Buffer - This depends on the polymerase and how much the cations have to be adjusted to achieve optimal polymerase activity.
Nucleotides - A final concentration of 200 mM of each dNTP works for most reactions. If you are making up dNTP from powdered stocks, you should add a pinch (a little off the tip of a spatula) of sodium bicarbonate to a few ml of 10 to 100 mM stock solutions and check that the pH is in the range of 7-8. Non-neutralized solutions of dNTP can rapidly hydrolyze.
Light mineral oil or paraffin wax - In the older PCR machines evaporation is a problem so mineral oil or wax is used to prevent evaporation. The newer PCR machines have lids that heat the entire tube and discourage condensation on the underside of the reaction tube which occurs in the older machines where the lid is unheated.
Subcloning Of PCR Products:
A typical PCR reaction yields a few micrograms of DNA. While this is a substantial amount of DNA one often needs to add to the PCR product sequences such as phage promoters to facilitate RNA synthesis or regulatory sequences so that the the protein product coded by the amplified DNA can be efficiently expressed. The simplest way to achieve these ends is to clone the DNA into an appropriate vector. Unfortunately, cloning PCR products is not as straightforward as it might appear. The reason for this is that some DNA polymerases used in PCR, notably Taq, possess a terminal transferase activity. i..e.. they can add non-template bases to the 3’ end of duplex DNA. Even the addition of a single base can prevent blunt-end cloning. There are two approaches to this problem which allow one to repair the ends. Either one can digest any single stranded overhangs with a 3’ to 5’ exonuclease such as T4 DNA polymerase, or one can fill in the oposite strand in a repair type reaction with a polymerase such as Klenow.
An alternative approach takes advantage of the overhangs. This approach is based upon the fact that the non-templated base is usually a single A residue. Therefore, if one has a vector containing a single overhanging T residue, one can clone the PCR fragment by a sticky end ligation into the vector. This procedure is called TA cloning. There are a number of ways of creating a T-tailed vector. One approach (D. Kovalic et al. Nucleic Acids Res. 19:4560 (1991), B. Schutte et al. Biotechniques 22:40 (1997)) is to obtain a vector which has an engineered site for XcmI in the polylinker. This enzyme leaves T overhangs. Other strategies for making TA vectors are given in Marchuk et al. Nucleic Acids Res. 19:1154 and Holton and Graham Nucleic Acid Res. 19:1156. A good source for information on making blunt end vectors is given in Costa and Weiner, Protocols For Cloning And Analysis Of Blunt-Ended PCR-Generated DNA Fragments, PCR Methods And Applications 3:S95-S106, Cold Spring Harbor Press. Because sometimes bases other than A are added (or no base at all), TA cloning is not a panacea for cloning PCR fragments. Furthermore, some enzymes with a high 3’ to 5’ exonuclease activity, e.g. Pfu, do not tend to add any extra bases. Finally, the extent to which extra bases are added can depend on the details of the PCR protocol. The use of an excessive number of cycles, or use of a final “extension cycle” of many minutes which is designed to make sure that all the PCR products are fully extended, will lead to the addition of more non-templated bases. If you use any sort of overhang vector, be sure to store the linearized vector lyophilized until ready for use and do not let it stay in solution for an extended period of time because the single stranded overhangs are extremely sensitive to any exonuclease contamination.
Finally, a completely different approach to cloning PCR products is to introduce restriction sites into the 5’ end of the PCR primers used for amplification. Following the PCR reaction, the products are digested with the appropriate enzymes and cloned into suitably cut vectors. One problem with this approach is that many restriction enzymes do not cut efficiently at the end of the DNA fragment. Therefore, workers usually include a GC “clamp” of at least 4 bases, 5’ to the restriction site, e.g. GCGC. This extension moves the restriction site away from the terminus to ensure efficient cleavage. The reason GC sequences are used is that GC is the most stable dinucleotide, ensuring the restriction enzyme will see a duplex terminus. The New England Biolabs catalogue contains a table showing the efficiency of cleavage near the terminus for a variety of restriction enzymes.
Sequencing of PCR Products. If a PCR product has been cloned into an appropriate sequencing vector (see above), sequencing the product is no different than for sequencing any other DNA molecule-except that the tendency of PCR to produce mutations and rearrangements of DNA must be carefully considered. For example, since clones are derived from a single DNA molecule if that molecule contains a PCR-derived mutation, then an incorrect sequence will be obtained. In addition, cloning PCR products is not without complications and even if everything works well, it takes additional time and materials. Hence a variety of protocols have been developed for direct sequencing of PCR products (B. Andersson and R. Gibbs in The Polymerase Chain Reaction K. B. Mullis et al. Birkhäuser Boston (1994) Chapter 7. F.M. Ausubel, Current Protocols in Molecular Biology, Unit 15.2, John Wiley (1995)). While it might not seem that sequencing a double stranded PCR product would be that much different from sequencing a double stranded covalently closed plasmid molecule, experience has shown that direct PCR sequencing is far trickier. For one thing, the crude PCR mix contains a large excess of primers and buffer salts that can interfere with sequencing. For this reason, sequencing with labeled nested primers internal to the primers used for amplification is often used. This strategy circumvents possible priming artifacts due to the failure to completely remove amplification primers since sequences primed from such primers will not be labeled.
The general problems that occur during direct PCR sequencing reflect the tendency of the complementary strand of the linear PCR product to reanneal and displace the sequencing primer and the impurities introduced into the sequencing reaction by the PCR step as noted above. The first cited reference by Andersson and Gibbs gives 13 (!) different methods that have been designed to overcome these problems. The journal Biotechniques has even collected 20 papers on PCR sequencing into a new book (The PCR Technique: DNA Sequencing Ed. J. Ellingboe and U.B. Gyllensten (1992)). At the present time we do not have sufficient experience with any of the methods to recommend one over the others. Not surprisingly, a variety of commercial kits are available to assist those with sufficient money to solve their PCR sequencing problems.
PROCEDURE 8.1: PREPARATION OF BACTERIAL AND MAMMALIAN DNA FOR PCR USING CHELEX
INTRODUCTION:
Chelex is a resin of styrene divinylbenzene copolymers containing iminodiacetate ions. These ions act to chelate polyvalent metal ions that might degrade the DNA during boiling. To isolate DNA with Chelex, the sample is boiled with the Chelex, and then the mixture is centrifuged. Then the DNA can be used immediately. This method is suitable for DNA isolation from blood, cells, hair roots, and bacterial cells. It is very quick and simple, a single prep takes approximately 45 minutes. However, the DNA that is isolated is single stranded and is, therefore, suitable for PCR, but it is not suitable for restriction enzyme analysis.
This technique works for many applications, but should be considered a crude preparation. If it works with minimal trouble-shooting, it will save you time processing samples. However, there are many template-primer combinations that will require more highly purified DNA in order to work.
PROCEDURE:
Use aerosol barrier tips for all subsequent pipetting steps.
1. For each sample add 200 ml of the 5% Chelex solution to a labeled 0.5 ml tube.
Note: Before removing the 200 ml of 5% Chelex for the extraction, be sure to vortex the solution so that the Chelex forms a homogenous suspension. Quickly, before the mixture can settle, aseptically transfer 200 ml of the Chelex to the 0.5 ml microcentrifuge tube. Use a 1.0 ml pipette tip for the transfer to avoid plugging the pipette tip with the resin.!
2a. For Cheek Scrape - Scrape the inside of the cheek with a toothpick and swirl the toothpick directly into the tube containing 200 ml of 5% Chelex.
2b. For Bacterial sample - Touch a toothpick to a bacterial colony and swirl the toothpick directly into the tube containing 200 ml of 5% Chelex.
Note: There have been reports that some toothpicks contain inhibitors that cause PCR reactions to fail. You may therefore wish to test each stock of toothpicks after they are first autoclaved. If you suspect toothpick poisoning you may wish to transfer colonies or plaques using either the end of a plastic pipette tip or plastic toothpicks.
2c. Place ~106 mammalian cells a tube containing 200 ml of 5% Chelex. You will have to optimize the number for your particular primers.
3. Incubate at 56oC for 15 to 30 minutes.
4. Vortex at high speed for 5 to 10 seconds. Keep your index finger on the cap of the tube during vortexing to prevent spilling any liquid!
5. Incubate in the DNA Thermal Cycler at 99 oC for 8 minutes.
6. Vortex at high speed for 5 to 10 seconds. Keep your index finger on the cap of the tube during vortexing to prevent spilling any liquid!
7. Spin in a microfuge tube for 2 to 3 minutes at 10,000 X g to 15,000 X g.
8. Remove supernatant to a fresh polypropylene tube. Be careful not to carry over any Chelex resin. If some Chelex resin is carried over, repeat the centrifugation step.
Note: If the resin is carried over to the amplification step it will inhibit the PCR.
9. Use 30 ml of the supernatant for 50 ml amplification as indicated in Protocol 8.2.
10. The remainder can be stored at 2oC to 8oC or frozen.
MATERIALS:
1. 5% Chelex
Compound Amount/ ml
H2O 95 ml
Chelex 5 g
2. 56oC Water bath
3. Microfuge and Microfuge Tubes
4. Toothpicks that will not inhibit PCR reaction
5. Biological Sample
PROTOCOL 8.2: GENERALIZED PCR REACTION
INTRODUCTION:
The following table gives the recipe for a PCR reaction. It is important to note that many variables affect the success of a given PCR procedure, and that it is often necessary to alter the procedure. Among the variables that can be changed are 1) primer sequence and concentration, 2) Mg++ concentration, 3) type of enzyme, 4) presence or absence of "specificity enhancers" such as DMSO, Formamide, or E. coli single stranded binding protein (Stratagene Perfect Match). This protocol is given assuming you wish to do a hot start procedure using paraffin wax. This helps prevent false priming that can occur as the sample is heating up on the first cycle of a normal PCR reaction.
Modifications to the protocol for a non-hot start PCR reaction using mineral oil is given in parentheses. If you are going to be adding the same volume of DNA to each tube, then you can combine the following reagents into a master mix and aliquot into individual tubes. The following table was set up assuming a 100 ml PCR Reaction.
Total ml Per Reaction ml
______ 77 ml H2O - volume of DNA template being added
______ 10.0 ml 10X PCR Buffer
______ 6.0 ml 25 mM MgCl2 (1.5 mM final)
______ 2.0 ml 10 mM dNTPs
______ 2.0 ml of forward primer (10 to 100 pmoles/100 ml rxn, typically 50-100 ng)
______ 2.0 ml of reverse primer (10 to 100 pmoles/100 ml rxn, typically 50-100 ng)
______ 1.0 ml of diluted Taq polymerase (2.5 U Taq).
Mix thoroughly
PROCEDURE:
Use aerosol barrier tips for all subsequent pipetting steps.
1. Check file parameters of the PCR machine. For example:
94oC, 1 min. Denaturation Temperature
50oC - 72oC, 1 min. The annealing temperature is dependent on the oligos being used
72oC, 2 min. Extension temperature
Note if annealing temperature is 72oC the annealing and extension steps can be combined into a single step.
1 cycle 72oC for 10 minutes, 4oC soak. This allows ample time for all the reactions to come to completion. (The soak step is optional and is not available on many machines)
2. It is often convenient to make a master mix containing all the PCR components except the DNA template to minimize pipetting steps and the chances of contamination.
Note 1: It is possible to do a modified hot start in some cases when you are not using a hot start polymerase. In this procedure you add all the components of the PCR reaction to a 0.5 ml microfuge tube on ice. Then start the PCR machine and when the temperature approaches 85oC quickly add the tubes to the thermocycler. This minimizes the amount of time that illegitimate priming can go on and often gives as good a reaction as a hot start using hot start polymerases.
Materials
1. Sterile H2O
2. 10X PCR Buffer - Depends on the polymerase being used
3. 10 mM dNTPs
4. Forward primer (50-100 ng/ml)
5. Reverse primer (50-100 ng/ml)
6. Heat stable DNA polymerase
7. Thermocyler
8. Mineral oil (Sigma M-3516) or Ampliwax Beads from Perkin Elmer.
9. Cells
PROTOCOL 8.3: AMPLIFICATION OF GENOMIC DNA
SOURCE: KIM, H.S. AND SMITHES, O. 1988. RECOMBINANT FRAGMENT ASSAY FOR GENE TARGETING BASED ON THE POLYMERASE CHAIN REACTION. NUCLEIC ACIDS RES. 16:8887-8903.
INTRODUCTION:
The following protocol gives a convenient method for lysing cells, extracting DNA, and performing a PCR reaction. The DNA extracted by this protocol is likely to vary in pH and Mg++ content and may require optimization. Stratagene sells a convenient kit for doing this optimization that is described in Protocol 8.5
PROCEDURES:
Use aerosol barrier tips for all subsequent pipetting steps.
Cell Lysis
1. Pellet up to a maximum of 1 X 106 cells for 30 sec. in microfuge. If more than 1 X 106 cells are used, the reagent volumes will have to be increased accordingly. For adherent cells, a trypsin treatment is generally used prior to this step to free the cells into suspension. Be careful to perform the trypsin step in a timely manner so that the cells do not remain in undiluted tyrpsin for a long period of time.
2. Aspirate media and wash cells with 500 ml of 1X PBS.
3. Pellet cells 30 sec. in microfuge at 14,000 rpm.
4. Aspirate PBS and resuspend cells in 500 ml of PBS. Repeat spin and aspiration steps.
5. Resuspend cells in 100 ml PBS and 200 ml sterile H2O.
6. Lyse cells by heating to 95oC for 10-15 minutes.
7. Allow the cells to cool briefly by setting at room temperature for 5 min. and add 10 ml of 10 mg/ml Proteinase K to each sample. Quick vortex to mix.
8. Incubate at 55oC for 1 hour.
9. Inactivate Proteinase K by heating to 95oC for 10 minutes.
10. Spin down condensation
11. Store at -20oC.
Use 10 ml for 50 ml PCR reaction. Note: In the subsequent PCR reactions you may have to carry out pH and Mg++ optimization to get the reaction to work.
MATERIALS:
1. Proteinase K (10 mg/ml)
· Dissolve proteinase K at 20 mg/ml in sterile deionized H2O.
· Aliquot and store at -20oC.
· The working concentration is 50 mg/ml.
3. 1X Phosphate Buffered Saline, pH 7.4 (PBS)
Compound Amount/ 1000 ml
Deionized H2O 800 ml
NaCl 8 g
KCl 0.2 g
Na2HPO4 1.44 g
KH2PO4 0.24 g
· Adjust pH to 7.4 with HCl.
· Add deionized H2O to 1 L.
· Dispense the solution into aliquots and sterilize them by autoclaving for 20 minutes at 15 lb/sq. in. on liquid cycle.
· Store at room temperature.
4. Waterbath at 95 oC
5. Plasticware
6. Microfuge
PROCEDURE 8.4: ELECTROPHORESIS IN NUSIEVE AGAROSE
INTRODUCTION:
This procedure is essentially identical to our standard method of DNA electrophoresis in agarose gels except that we use a special grade of agarose which is particularly useful for resolving DNA fragments <1 kb. We use a 3%:1% (4% Total) Nusieve:SeaKem Gold agarose gel (1X TBE). The NuSieve gel approaches the resolution of acrylamide and is good for visualizing small DNA fragments. By itself, NuSieve is rather brittle and hard to work with. The addition of 1% SeaKem Gold agarose gives the gel much greater strength without loss of resolution. The one caution is to not run the gel too hot (greater than 50V) because NuSieve will melt at higher temperatures. Note that BioWhittaker (formally FMC) now sells a mixture of Nusieve:Sea Kem Gold Agarose 3:1 obviating the need to make one's own mixture.
PROCEDURE:
1. Weigh out 3 g NuSieve Agarose and 1 g SeaKem Gold agarose, add 100 ml 1X TAE, boil briefly in the microwave, then swirl gently until all the particles are dissolved.
2. Casting the Gel
A. Place clean 2 X 3 inch plate in the bottom of the casting tray.
B. Draw up molten agarose in Pasteur pipette and run a bead of agarose around the edge of the casting tray so that the agarose cannot leak out of the tray. Let harden a few minutes.
C. Carefully pipette cooled (55oC) agarose into tray (12-15 ml) in 1X TBE until the agarose comes to the edge of the casting tray.
D. Gently place a comb into the slots of the casting tray to form the wells of the gel.
E. When the gel has hardened (10 minutes), flood the surface with buffer and carefully remove the comb, one end first. If the gel is not fully hardened, it may help to briefly chill it in the refrigerator prior to removing the comb, in order to prevent partial collapse of the sample wells. These wells should hold about 10-20 ml depending on the thickness of the gel.
F. Remove gel from casting tray by gently pushing up through the hole in the bottom and place in electrophoresis apparatus.
G. Use 1X TAE buffer (preferably cold) as a running buffer; fill the submarine gel chamber to just over the top surface of the gel.
Alternatively use precast gels from FMC.
3. Loading the gel
Use aerosol barrier tips for all subsequent pipetting steps.
A. Remove 9 ml of PCR (100 - 400 ng) reaction and add 1 ml of 10X loading dye.
B. Load all 10 ml of digest per well.
C. In a mini-gel horizontal gel apparatus, electrophorese at 50 to 75 volts until the Bromophenol blue dye front is 1/2 - 2/3 way down the gel (Note: do not run Nuseive gels at voltages above 100V because they can melt). The Bromophenol blue dye front moves about where a 300 bp piece of DNA would run.
D. Stain gel in 0.1 mg/ml EtBr for at least 15 minutes; photograph the gel.
NOTE: Agarose gels may be stored several days at 4oC wrapped in Saran Wrap. The BioWhittaker (Formally FMC) catalogue presents a wealth of useful information concerning the different types of agaroses available and their specific uses.
Ethidium bromide is a mutagen. ALWAYS wear gloves and dispose of in waste into container so that it can be detoxified at a later time (see the procedure in the Safety section of the manual).
Note that the concentration of EtBr suggested in this protocol is considerably lower than that given in many other protocols. However it produces very clear staining with minimum background fluorescence. Staining for up to 1 hour gives somewhat higher sensitivity. Very long staining times, e.g. overnight can lead to blurring and even loss of lower molecular weight bands.
MATERIALS:
1. Casting tray and plates.
2. Agarose and casting trays.
3. 5 X TBE
Compound Amount/ 1000 ml
Tris base 54 g
Boric Acid 27.5 g
0.5 M EDTA, pH 8.0 20 ml
Deionized H2O to volume
· Mix, dispense into aliquots, and sterilize by autoclaving.
· A precipitate forms when concentrated solutions of TBE are stored for long periods of time. To avoid problems, store the 5X solution in glass bottles at room temperature and discard any batches that develop a precipitate.
· TBE buffer can be reused many times.
4. 50 X TAE
Compound Amount/ 1000 ml
Tris base 242 g
Glacial Acetic Acid 57.1 ml
0.5 M EDTA, pH 8.0 100 ml
Deionized H2O to volume
· Mix, dispense into aliquots, and sterilize by autoclaving.
5. Ethidium Bromide (10 mg/ml, FW=394.3)
Compound Amount/10 ml
Deionized H2O 10 ml
Ethidium Bromide 0.1 g
· Carefully add the ethidium bromide to a 15 ml conical centrifuge tube and add the water.
· Cap the solution tightly and rinse off the outside of the tube.
· Vortex periodically over for several hours to ensure that the dye has dissolved.
· Wrap the tube in aluminum foil or transfer the solution to a dark bottle and store at room temperature.
Caution: Ethidium bromide is a powerful mutagen and is moderately toxic. Gloves should be worn at all times. Wear a mask when weighing the solution out.
6. Electrophoresis box and power supply.
PROTOCOL 8.5: PCR AMPLIFICATION OF CLONED INSERTS FROM LAMBDA PHAGE AND BACTERIAL COLONIES
Introduction:
A common way to amplify a cloned insert is to use oligonucleotide primers specific to vector sequences just flanking the cloned insert. Commonly these primer pairs are M13 Forward and M13 Reverse, T7 and T3 or T7 and SP6. Large amounts of DNA can be generated this way without having to grow large amounts of lambda or plasmid DNA. PCR can also be used to check the size of specific clones. In the case of cDNA libraries, a number of clones can be combined in one PCR reaction and the average insert size of the library estimated. This specific protocol has been used to amplify inserts from lambda clones. Note however that some types of wooden toothpicks contain materials that can inhibit the PCR reaction. Therefore, it is recommended to use glass pipettes, plastic toothpicks or disposable pipette tips for picking plaques or colonies.
PROCEDURE:
Use aerosol barrier tips for all subsequent pipetting steps.
For Bacterial Colonies
1. Take plastic pipette tip and lift a colony from the plate taking as little agar as possible.
2. Twirl pipette tip in 50 ml of water in microfuge tube to remove bacteria.
Note: If you the bacteria used to make the PCR template are needed for additional work, take the tip and streak a gridded selection plate as described in toothpick procedure in Protocol 2.4
3. Boil sample 10 minutes in screw top tube.
4. Spin briefly in microfuge tube to settle condensate and cell debris.
5. Place on ice or freeze until ready for use in PCR reaction.
6. Continue with PCR below.
For lambda phage
1. Randomly pick 2 colorless plaques from your high dilution titer plate using the small end of a Pasteur pipette.
2. Blow out end of plug into a 1.5 ml microfuge tube containing 200 ml water.
3. Incubate tubes at room temperature for >30 min. then freeze at -20oC until ready for PCR. (Warm up 15 min. prior to use).
4. Continue with PCR below.
Assemble PCR
1. Assemble PCR mix. For each 20 ml reaction add the following components:
2 ml 10X PCR Buffer
3.2 ml 1.25 mM dNTPs
2 ml of forward primer ( e.g. T7 primer (50 ng/ml)
2 ml of reverse primer (e.g. T3 primer (50 ng/ml)
0.3 ml Hot Tub (3 U/ml - Amersham) or 0.1 ml Taq DNA polymerase (5 U/ml)
Mix thoroughly
Immediately before performing the PCR reaction make a master mix containing all these reagents. Multiply the above volumes by the number of samples, in this case 3, to estimate the amount of each reagent needed. You should usually allow for on extra reaction to be sure you do not run out of master mix due to pipetting errors.
NOTE: To prevent contamination use the plugged tips that are provided.
3. Aliquot 9.5 ml master mix into sterile 0.5 ml microfuge tubes.
4. Add 10.5 ml sample DNA to 2 tubes tube and 10 ml H2O to a third tube which serves as the no DNA control. Mix and spin 2 sec.
5. Overlay sample with 15 ml of mineral oil.
6. Place tubes in PCR machine and run appropriate file.
7. File Parameters are dependent on the length of the insert being amplified and the annealing temperature of the primers :
E.g. the following parameters work for the standard T7 and T3 oligonucleotides for most inserts less than 3 kb.
94oC, 1 min.
55oC, 1 min.
72oC, 3 min.
30 cycles: approximately 3 hours 40 min.
1 cycle 72oC for 10 minutes, 4oC soak.
8. Remove samples; place on ice or store in refrigerator.
11. Remove 10 ml and load on a 1.2% agarose gel (1X TBE). Run at 50 to 100 volts until the bromophenol blue dye front is 1/2 - 2/3 way down the gel.
MATERIALS:
The same as in Protocols 8.3 and 8.4
PROTOCOL 8.6: PRIMER DESALTING
Reference: B Freie and S.H. Larsen Biotechniques 10:420 (1991)
INTRODUCTION:
Most modern oligonucleotide synthesizers generate relatively few incorrect chains when the oligonucleotides are less than approximately 25 bp. Thus for most PCR reactions it is unnecessary to purify the oligonucleotides on a gel and isolate the fragment corresponding to the proper sized primer. The dried reaction products you typically receive from the synthesizer facility, however, may contain a lot of salts and organic contaminants that can interfere with PCR reactions. Thus it is recommended that you desalt your oligos before use or have this done by the sequencing facility. The following is an easy, inexpensive procedure for desalting and cleaning up oligonucleotides using Waters C18 SepPak columns. These columns have a capacity to retain large amounts of oligonucleotides. Therefore, even a 1.0 mM reaction can be purified using these columns.
PROCEDURE:
1. Wash Sep-Pak column with 5 ml methanol followed with 5 ml of water. You push these through slowly using a 5 or 10 ml syringe.
2. Load sample in 0.3 - 1.0 M NaCl. Typically the oligo is diluted to 1.0 ml with 0.5 M NaCl.
3. Wash column with 2 ml water. Again use slow steady push.
4. Elute oligo with three 1 ml washes of 60% methanol/water.
5. Read O.D.260 of the three fractions. Greater than 90% of the oligo should be in the first fraction.
6. Combine the fractions with significant O.D. and dry in a Speed Vac. You should allow a half day or more for the oligonucleotide to completely dry.
7. Resuspend oligo in sterile water or TE, pH 8.0. For a 1.0 mM synthesis this is typically 1.0 ml. Measure O.D.260 and determine concentration by 32 X O.D. X dilution factor = mg/ml.
MATERIALS:
1. Sep-Pak Column
2. 5 M NaCl (FW=58.44)
Compound Amount/ 1000 mll
Deionized H2O 800 ml
NaCl 292.2 g
· Dissolve and adjust the volume to 1 L with deionized H2O.
· Dispense into aliquots and sterilze by autoclaving.
3. 60% Methanol
Compound Amount/ 500 ml
Methanol 300 ml
H2O 500 ml
4. Spectrophotometer
5. Lyopholyzer
6. TE, pH 8.0
Final Concentration Stock Solution Amount/ 100 ml Amount/ 500 ml
Deionized H2O 90 ml 400 ml
10 mM Tris Cl 1 M, pH 8.0 800 ml 4 ml
1 mM EDTA 0.5 M, pH 8.0 80 ml 400 ml
Deionized H2O to volume
· Mix and dispense into aliquots. Sterilize by autoclaving. Store at room temperature.
·
7. Sterile H2O
PROTOCOL 8.7: OPTIMIZING PCR REACTIONS
INTRODUCTION:
As described in the introduction many factors influence the fidelity of a PCR reaction, particularly when the source of the DNA is from a cell lysate or cDNA library where pH and Mg++ are likely to be variable. Oftentimes the easiest way to optimize a PCR reaction is to take an empirical approach where aliquots of the target DNA is amplified under a number of different conditions to determine which one works best. Once the system is optimized, it is often possible to use those conditions on a consistent basis.
The two most commonly varied conditions are pH and Mg++ which dramatically effect how polymerases function. A number of other modifiers, such as DMSO and single strand binding protein, can also be used to enhance yield and fidelity. Perkin Elmer publication Amplifications is good source of information on the factors affecting PCR reactions and the latest developments in PCR protocols.
The following protocol is from the Strategene PCR optimization kit. This protocol begins by doing 12 separate PCR reactions covering a range pH, Mg++, and KCl concentrations. If one or more of these conditions looks promising, other modifiers such as DMSO are used to fine-tune the reaction.
PROTOCOL:
A. Determining the Optimal Buffer Concentration. Use aerosol barrier tips for all subsequent pipetting steps.
1. Label 12 sterile microcentrifuge tubes with the number 1 through 12. Add 5 ml of each of the 12 Opti-Prime 10X buffers to its corresponding microcentrifuge tube.
2. Place a separate sterile microcentrifuge tube on ice and add the following components in order:
NOTE: The following components will make a reaction mixture sufficient for ~12.5 PCR amplification reactions each of 50 ml. Only 12 PCR amplification reactions will be performed, but the extra amount will ensure a sufficient volume of the buffer. Add the components in the order listed below.
Master Mix (12.5 reactions)
565-(X+Y + 50 ml) Deionized H2O sufficient to bring the total volume to ~565 ml
12.5 ml 10 mM dNTPs (2.5 mM each dNTP)
12.5 ml Master Mix 50X buffer (final concentration of 400 mM Tris-HCl, pH 8.0 and 5 nM EDTA)
12.5 ml 3’ Primer (Assuming 100 ng in 1 ml)
12.5 ml 5’ Primer (Assuming 100 ng in 1 ml)
X ml DNA (2500 ng genomic DNA in master mix so that there will be 200 ng genomic DNA in each final reaction)
Y ml 30 U Taq DNA Polymerase
N.B. Addition of the Master Mix buffer helps protects the DNA polymerase until the Master Mix is added to the various 10X Optiprime buffers.
3. Add 45 ml of the above reaction mixture to each of the 12 tubes from step number 1.
Note: if you are using a PCR machine without a heated lid you would have to layer 25 ml of mineral oil over each of the 12 microcentrifuge tubes to seal the reactions in order to prevent excess evaporation.
4. Place the microcentrifuge tubes in the thermocycler and initiate the amplification reactions. The table below lists a sample time schedule for a 1000-bp PCR product:
PCR steps Temperature Duration
a. Denature 94oC 3 minutes
b. Anneal 50oC 2 minutes
c. Repeat the following cycles 20-30 times
i. Extend 72oC 1.5 minutes
ii. Denature 94oC 1 minute
iii. Anneal 50oC 1 minute
d. Extend 72oC 8 minutes
5. Load 10 ml of each of the 12 PCR samples onto an agarose or 6% (w/v) acrylamide gel or 2% 3:1 Nusieve agarose gel. Be sure to add 1 ml of 10X loading dye to the samples before loading. Electrophorese, stain and evaluate the PCR products for their correct size, for the desired PCR product yield and for the amount of nonspecific background amplification products present.
6. From the results tabulated in step 5 above, the best PCR buffer for this DNA template and primer set can be determined. If desired, choose the best buffer (s) and further optimize by the addition of adjuncts (see Optimal Adjunct Determination).
Optimal Adjunct Determination
1. Select the appropriate Opti-Prime 10x buffer as determined from the optimal buffer assay described in the previous section (see Optimal Buffer Determination). Prepare a reaction mixture of all the components (excluding the adjuncts, which will be added separately) as indicated below.
Place a sterile microcentrifuge tube on ice and add the following components in order:
NOTE: The following components will make a reaction mixture sufficient for ~7.5 PCR amplification reactions in order to ensure an adequate volume of the buffer. In this case, however, the optimal Opti-Prime 10x buffer will be used as determined in the optimal buffer assay. (see Optimal Buffer Determination).
A. 340 - (X+Y + 60) ml Deionized H2O sufficient to bring the total volume to ~340 ml
B. 37.5 ml Optimal Opti-Prime 10x buffer (see Optimal Buffer Determination).
C. 7.5 ml dNTPs 10mM dNTPs (2.5 mM each dNTP)
D. 7.5 ml 3’ Primer (Assuming 100 ng in 1 ml)
E. 7.5 ml 5’ Primer (Assuming 100 ng in 1 ml)
E. X ml 1.5 mg of genomic DNA template (200 ng/reaction) or 750 ng of the plasmid DNA template (100 ng/reaction)
F. Y ml 18.0 U Taq DNA polymerase
2. Label seven sterile microcentrifuge tubes with the number 1 through 7. Add the adjuncts to each of the seven microcentrifuge tubes as indicated in the table below:
Tube Number Adjunct Final Concentration
1 5.0 ml of formamide 5%
2 2.5 ml of DMSO 5%
3 7.5 ml of glycerol 15%
4 1.0 ml of 750 mM (NH4)2SO4 15 mM
5 3.0 ml of 1.5 mg/ml BSA 10 mg/ml
6 0.5 U of Perfect MatchâDNA polymerase enhancer (genomic templates) or 0.05 U of Perfect MatchâDNA polymerase enhancer (plasmid templates)
7 5.0 ml of Sterile dH20
3. Add 45 ml of the reaction mixture from step 1 to each of these seven microcentrifuge tubes. Overlay 25 ml of mineral oil onto each reaction and seal the microcentrifuge tubes.
NOTE: The final volume of each amplification reaction is not exactly 50 ml. This slight variance is insignificant and will not affect the final results.
4. Place the seven microcentrifuge tubes in the thermocycler and cycle according to the calculated parameters. (see Preprotocol Considerations for the suggested guidelines).
5. Load 10 ml of each of the seven PCR samples onto an agarose or 6% (w/v) acrylamide gel or 4% 3:1 Nusieve agarose gel. Electrophorese, stain and evaluate the PCR products for their correct size, for the desired PCR product yield and for the amount of nonspecific background amplification products present.
6. At this point, the optimal Opti-Prime 10x buffer and the appropriate adjunct has been determined for the particular primer-template set.
MATERIALS:
1. 10X Opti-Prime Buffers
Opti-Prime 10x Buffer #1 Opti-Prime Buffer 10x #2
100 mM Tris-HCl (pH 8.3) 100 mM Tris-HCl (pH 8.3)
15 mM MgCl2 15mM MgCl2
250 mM KCl 750 mM KCl
Opti-Prime 10x Buffer #3 Opti-Prime Buffer 10x #4
100 mM Tris-HCl (pH 8.3) 100 mM Tris-HCl (pH 8.3)
35 mM MgCl2 35mM MgCl2
250 mM KCl 750 mM KCl
Opti-Prime 10x Buffer #5 Opti-Prime Buffer 10x #6
100 mM Tris-HCl (pH 8.8) 100 mM Tris-HCl (pH 8.8)
15 mM MgCl2 15mM MgCl2
250 mM KCl 750 mM KCl
Opti-Prime 10x Buffer #7 Opti-Prime Buffer 10x #8
100 mM Tris-HCl (pH 8.8) 100 mM Tris-HCl (pH 8.8)
35 mM MgCl2 35mM MgCl2
250 mM KCl 750 mM KCl
Opti-Prime 10x Buffer #9 Opti-Prime Buffer 10x #10
100 mM Tris-HCl (pH 9.2) 100 mM Tris-HCl (pH 9.2)
15 mM MgCl2 15mM MgCl2
250 mM KCl 750 mM KCl
Opti-Prime 10x Buffer #11 Opti-Prime Buffer 10x #12
100 mM Tris-HCl (pH 9.2) 100 mM Tris-HCl (pH 9.2)
35 mM MgCl2 35mM MgCl2
250 mM KCl 750 mM KCl
2. Master Mix 50X Buffer
Final concentration of 400 mM Tris-HCl (pH 8.0) and 5 nM EDTA
3. DNTP Stocks (10 mM)
4. Thermostable polymerase, e.g. Taq and thermocycler
PROTOCOL 8.8 REVERSE TRANSCRIPTION PCR
SOURCE: Gibco/BRL Technical Sheet
INTRODUCTION:
This procedure is slightly longer than some as it uses a heating step to denature the RNA prior to reverse transcription. Presumably this step helps to improve the ability of the reverse transcriptase (RT) to transcribe through regions of secondary structure. The RT used here is a genetically engineered mutant of the Moloney Leukemia Virus RT in which the RNase H activity of the enzyme has been removed by point mutations which improves the ability of the enzyme to generate full length transcripts.
PROTOCOL:
1. Mix:
1 ml of oligo dT 12-18 (500 mg/ml)
1-5 mg of total RNA
Sterile water to 12 ml
Heat to 70oC for 10 minutes and quick chill on ice. Spin to collect.
2. Add to the tube
4 ml of 5X First Strand Buffer
2 ml of 0.1 M DTT
1 ml of dNTP (10 mM each)
Mix gently. Incubate at 42 oC for 2 min. Add 1 ml of Superscript II RT (200U/ml). Pipette gently to mix. Incubate at 42 oC for 50 min. Inactivate the enzyme by heating for 15 min. at 70oC. The cDNA is now ready for PCR. Do not use more than 10% of this reaction in a 100 ml PCR reaction.
MATERIALS:
1. 5X First Strand Buffer:
250 mM Tris-HCl (pH 8.3 at room temperature)
375 mM KCl
15 mM MgCl2
2. 0.1 M Dithiothreitol
Store both solutions at -20oC. Thaw them at room temperature just prior to use and refreeze immediately.
PROTOCOL 8.9: SITE DIRECTED MUTAGENESIS BY INVERSE PCR
SOURCE: Stratagene QuikChangeTM Site-Directed Mutagenesis Kit
INTRODUCTION:
The following protocol is a general method for site-directed mutagenesis. It has several advantages over alternative protocols. It does not require 1) the use of special vectors, 2) it only requires the use of two primers, and 3) it uses only a single PCR reaction. The only disadvantage is that it requires PCR amplification of relatively large templates, which allows for errors outside the region of the template one desires to mutagenize. In addition, long PCR can be problematic and often requires the use of polymerase mixtures where one of the enzymes has a 3' to 5' exonuclease error checking activity.
The protocol works as follows. Two complementary primers are synthesized, each of which carries the mutation one wishes to introduce. These primers are designed so that following annealing to a covalently closed double-stranded plasmid containing the region one wishes to mutagenize, PCR will proceed away from the region being mutagenized in opposite directions (see diagram below). Next one carries out a series of PCR amplifications to synthesize new stands of the plasmid which contain the mutation. One of the most elegant aspects of this protocol is mechanism for removing the unmutagenized parental template. Plasmids grown in most E. coli strains are methylated on A residues of the sequence GAmetTC. Such DNA is susceptible to cutting by the restriction enzyme Dpn I. DNA synthesized in vitro normally does not contain such methylated residues and is therefore resistant to Dpn I digestion. Hence following PCR, a brief treatment with Dpn I serves to eliminate the parental template strands while leaving the mutagenized daughter strands. Following Dpn I digestion, the amplification mix can be use to directly transform highly competent E. coli cells. Following DNA uptake, ligation of the nicked of the nicked circular molecules occurs in vivo. A majority of the resulting transformants usually contains the desired mutation.
The following abbreviated protocol does not include several details on primer design, storage of materials, controls etc. that are provided in the Stratagene™ kit. Stratagene states in the kit instructions that the primers have to be highly purified, but this is probably not essential for many purposes. Nonetheless it is probably prudent to be sure that your primers are at least desalted when they are purchased. If they are not, then you will need to desalt them using either alcohol precipitation or reverse phase chromatography on C18 columns (Protocol 8.6).
It should be noted that the PCR reaction used in inverse PCR mutagenesis leads to a linear, not an exponential amplification. The reason is the 3' ends of the amplification primers are aligned in opposite directions in inverse PCR mutagenesis. Because DNA polymerases add bases only in a 5' to 3' direction, any primer that binds to the end of a newly synthesized linear strand will have no template from which to copy a new strand (see diagram below). Hence only the two parental plasmid strands will be copied.
3' Daughter Strand W 5' 3' < No Template For PCR Reverse Primer
A similar diagram would apply to daughter strand C. Thus neither daughter strand can be used as a template for further PCR, but they can anneal to each other to form a doubly nicked circle capable of transformation at low effeciency.
The success of inverse PCR mutagenesis is therefore very dependent on the substrate concentration. Too little template DNA, and not enough product is formed during the 12 to 20 cycles of amplification. When too much template DNA is employed, the primers can anneal non-specifically to alternate sites on the template leading to abortive PCR products that amplify preferentially over the template strands. One can often see these smaller Dpn I resistant PCR fragments even in successful template amplifications. Hence, in setting up the procedure, a range of template concentrations should be tested at a fixed primer concentration to empirically determine the template concentration that yields the greatest amount of fully replicated template. A typical range of starting template concentrations would be 10, 20, 40 and 100 ng per 50 ml PCR reaction.
PROTOCOL:
1. Prepare four different DNA template concentrations in the range of 5 to 50 ng / ml. Often 10 to 20 ng/ ml works best.
2. Make a master for 4.5 - 50 ml reactions.
173.1 - ml of Sterile water
22.5 ml of 10 x Reaction Buffer
5.6 ml of Primer 1 (100 ng/ml)
5.6 ml of Primer 2 (100 ng/ml)
4.5 ml of dNTP mix (10 mM in each dNTP*.)
5.5 ml of Pfu Turbo Polymerase (2.5 U/ml)
*Stratagene suggests not using material that has been subject to multiple freeze/thaw cycles.
3. Aliquot 48 ml of Master Mix into 4 PCR tubes.
4. Add 2 ml of the appropriate template concentration to each tube.
5. Cycle the reaction using the cycling parameters outlined in Table I below.
(N.B. The time of 2 min/kb of plasmid length given in the Table may be minimal. Stratagene actually recommends 12 minutes for a 4.5 kb plasmid. Also the number of cycles given may be minimal, and if a visible yield of product is not seen, one may wish to use a higher number of cycles. Do not use more cycles than necessary however as this will increase the probability of mutations introduced by the polymerase outside the region of interest)
TABLE I Cycling Parameters for the QuikChange Site-Directed Mutagenesis Method
Segment Cycles Temperature Time
1 1 95°C 30 seconds
2 12–18 95°C 30 seconds
55°C 1 minute
68°C 2 minutes/kb of plasmid length
6. Following completion of the reaction run 10 ml of the reaction mix on an agarose gel.
Stratagene suggests a 1% gel, but the actual concentration should be adjusted so that optimum resolution is obtained between the input and product DNAs which will make it easier to see if the PCR has been successful.
If you do not see any product you can precede with the Dpn I digestion step in any case, but you will have to ethanol precipitate the entire amplification mix in order to have a chance of obtaining colonies. It may be easier to optimize the PCR reaction instead.
7. Add 1 ml of Dpn I (10 U/ml).
This is an important point. If the mutation frequency is too low, it may be because of inadequate digestion of the parental template by Dpn I. In that case, increase the amount of Dpn I and/or extend the time of digestion.
6. Gently pipette up and down to thoroughly mix the reaction. Spin for a few seconds to collect the material and incubate for 1 h at 37oC.
7. Following digestion transform competent cells using either chemical transformation (or alternatively electroporation).
One needs high transformation frequencies to make the procedure effective. A procedure for XL-Blue Supercompetent cells (> 1 x 108 cfu/mg) follows:
8. Prechill Falcon 2059 tubes in an ice bath and add 1-4 ml of the Dpn I digested DNA to the tubes. Then thaw the competent cells on ice, mix gently and add 50 ml of cells to the tubes. Mix by swirling the tubes.
9. Incubate 30 minutes on ice. Mix by swirling occasionally.
10. Heat shock the cells by incubating for 45” at 42oC. Then immediately return the cells to the ice bath for 2 minutes.
11. Add 0.5 ml of NZY+Broth (prewarmed to 42oC) and incubate at 37oC for 1 h with shaking.
12. Plate all of the 0.5 ml on a fairly dry LB plate with appropriate antibiotic selection
13. Incubate at 37oC for at least 16 hours, but do not prolong the incubation longer than necessary to pick colonies to avoid the growth of satellite colonies.
14. Pick well-isolated colonies into LB with appropriate antibiotic selection and grow 5 ml cultures overnight for restriction enzyme digestions and/or sequencing.
MATERIALS:
1. LB Agar (per Liter)
10 g of NaCl
10 g of tryptone
5 g of yeast extract
20 g of agar
Add deionized H2 O to a final volume of
1 liter
Adjust pH to 7.5 with 5 N NaOH
Autoclave
Pour into Petri dishes (~25 ml/100-mm plate)
2. LB–Ampicillin Agar (per Liter) (One can use carbenicillin for reduced chances satellite colony formation)
1 liter of LB agar
Autoclave
Cool to 55°C
Add 50 mg of filter-sterilized ampicillin
Pour into Petri dishes (~25 ml/100-mm plate)
3. NZY+ Broth (per Liter)
10 g of NZ amine (casein hydrolysate)
5 g of yeast extract
5 g of NaCl
Adjust to pH 7.5 using NaOH
Autoclave
Add the following supplement prior to use
12.5 ml of 1 M MgCl2 and 12.5 ml of
1 M MgSO4
10 ml of a 2 M filter-sterilized
glucose solution or 20 ml of
20% (w/v) glucose
Filter sterilize
4. 10× Reaction Buffer
100 mM KCl
100 mM(NH4 )2 SO4
200 mM Tris-HCl (pH 8.8)
20 mM MgSO4
1% Triton ® X-100
1 mg/ml nuclease-free bovine serum
albumin (BSA) ‡