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 Two somewhat out-of-date texts which discuss a wide variety of PCR techniques are: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.

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.

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 93^oC to 94^oC so that the strands come apart. In the second step, the reaction is cooled to a lower temperature, typically 50^oC to 72^oC, such that the primers anneal selectively to the regions which flank the target site to be amplified. Lastly, the reaction is heated to 72^oC to 75^oC 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 70^oC. 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 90^oC 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 56^oC 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 2^oC to 8^oC or frozen.

MATERIALS:

1. 5% Chelex

Compound Amount/ ml

H₂O 95 ml

Chelex 5 g

2. 56^oC 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:

94^oC, 1 min. Denaturation Temperature

50^oC - 72^oC, 1 min. The annealing temperature is dependent on the oligos being used

72^oC, 2 min. Extension temperature

Note if annealing temperature is 72^oC the annealing and extension steps can be combined into a single step.

1 cycle 72^oC for 10 minutes, 4^oC 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 85^oC 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 95^oC 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 55^oC for 1 hour.

9. Inactivate Proteinase K by heating to 95^oC for 10 minutes.

10. Spin down condensation

11. Store at -20^oC.

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 -20^oC.

· 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 (55^oC) 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 4^oC 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