PCR仪

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I. Denature
  93-94 degrees C 1.5 minutes
  II. Anneal 50-65 degrees C 2 minutes
  III. Polymerize 72 degrees C 2 minutes
  Strategies for optimizing PCR reactions are at the end of the protocol.
  7.After completion of the PCR reaction, remove the tubes from the temperature block and wipe the outside free of excess oil before placing in an eppendorf rack.
  8.Add 2.0 祃 of 5X Ficoll stop dye directly into the aqueous phase "bubble" at the bottom of each tube, and then add 100 祃 of chloroform:isoamyl alcohol (24:1) to each tube, shake well, and spin briefly.
  9.Carefully remove only the aqueous "bubble" with a P20 pipetman set to 7-8 祃 by placing the pipet tip against the bubble and slowly drawing it in. Each sample should then be placed in a separate clean eppendorf tube before loading onto the polyacrylamide gel.
  10.The reaction products are conveniently separated according to size by electrophoresis through a 10% polyacrylamide "Mighty-small II" gel at 110 V for 2-2.5 hours, and visualized after staining the gel with ethidium bromide.
ADDITIONAL INFORMATION:
Preparation of Oligonucleotides:
Oligonucleotide primers are synthesized using an automated machine (we currently order primers through the Center for Genetics in Medicine) and are received in a glass vial in an ammomium solution. It is convenient to remove about one half of the total volume for each oligonucleotide and divide this volume further into two 1.5 ml eppendorf tubes (the remainder of the ammonium stock solution is stored at 4 degrees
C). The oligonucleotides must be prepared as detailed below before use in PCR reactions:
  1.Incubate each sample in a heating block at 55 degrees C overnight and then dry in a rotary vacuum concentrator for 4-6 hours. (Warning: a cold trap should be used when drying the samples to absorb the ammonia)
  2.Resuspend each oligonucleotide (from both eppendorfs) in a total of 500 祃 of TE.
  3.Make a 1:200 dilution by diluting 5 祃 of each oligonucleotidewith 1.0 ml of TE and measure absorbance of UV light in a spectrophotometer at 260 and 280 nm. The concentration of the stock of resuspended oligonucleotide can then be calculated:
  A260 of 1.0 = 35 礸/祃 for DNA oligonucleotides. If A260 = 0.203 for a oligomer of 21 nucleotides, then 0.203 x 35 x 200 (dilution) = 1421 礸/ml (original solution),or 1.421 x 106 礸/L 21 (#nucleotides) x 330 礸/祄ol = 6930 礸/祄ol 1.421 x 106 礸/L = 205 祄ol/L (礛) ---------------- 6930 礸/祄ol
  4.Make 50 礛 solutions in TE of each oligonucleotide for subsequent use in PCR reactions.
Strategies for optimizing the efficiency of PCR reactions:
  The conditions required for generation of a specific, essentially unique product (single strong band) will nearly always need to be optimized empirically. In particular, the annealing temperature is important in determining the specificity of the reaction (that is to say, at lower temperatures the primers may anneal to similar irrelevant sequences elsewhere in the genome and prime these, resulting in the formation of multiple products). In general, higher annealing temperatures result in more stringent conditions for primer annealing and more specific products. A good place to start is with a low annealing temperature around 50-55 degrees C, with optimization by testing at 3-5 degrees C increments until maximum specificity is reached. Theoretically, oligonucleotide primers with a high GC content may require a very high annealing temperature to maximize specificity. While this is a good rule of thumb, the optimum temperature may not correspond well to this estimate. Occasionally, specifity will reach a maximum at a certain temperature and at higher annealing temperatures, multiple new products or no products at all will be generated. Although annealing temperature is perhaps the easiest variable to change, specificity may also be increased by reducing the concentration of primers or Taq polymerase, minimizing the times allowed for annealing and extension, or reducing the free Mg++ concentration. An optimum of Mg++ concentration usually exists in the 1-10 mM range. Too low Mg++ concentration may result in no
products and an excess may result in a variety of unwanted products.
Pouring and Running Polyacrylamide Gels using the Hoefer SE-250
"Mighty-small II" gel electrophoresis unit: (Simplified instructions are provided below, for detailed instructions, refer to the Hoefer manual). Multiple identical polyacrylamide gels can be pre-cast in the supplied SE 275 multiple gel caster. Acrylamide is a neurotoxin and should be handled with caution. Wear gloves at all times when handling acrylamide and be careful to avoid spills.
  1.Clean the multiple gel caster and place flat on the bench top in front of you. Place the rubber gasket in its groove without stretching it and lubricate with a thin layer of the Cello-seal provided by Hoefer.
  2.Build the gel casting units by carefully placing and seating components in the following order from the bottom up: waxed paper, notched alumina plate, T-shaped spacers (0.75 or1.0 mm), glass plate, waxed paper, etc. Approximately 5 complete 0.75 mm gels can be cast at one time with one or two additional glass plates needed to fill extra space.
  3.Place the top cover on the multiple gel caster and apply red spring clamps to side grooves, ensuring adequate sealing. Be sure that the port at the bottom of the front plate has a small piece of rubber tubing on it and is clamped off.
  4.Mix the ingredients for 50 ml of acrylamide (minus the TEMED) in a clean beaker, as detailed in the recipe below for a 10% polyacrylamide gel. Add the TEMED with thorough mixing just before pouring the gels.
  5.Carefully pour the acrylamide evenly into the gel casting units in the multiple gel caster until the multiple gel caster is almost overflowing.
  6.Insert the appropriate sized comb (0.75mm for 0.75 mm spacers) into each gel casting unit, and allow the acrylamide to polymerize for at least 1 hour. After complete polymerization, the gels may be wrapped in cellophane and stored at 4 degrees C.
Solutions:
  40% Acrylamide/ 2% bis stock
  acrylamide: 38 g
  N,N'-methylene bisacrylamide 2 g
  dH20: to 100 ml
  Mini-gel ("Mighty-small II") 10% polyacrylamide (50 ml volume)
  12.5 ml 40% Acrylamide/ 2% bis stock
  25 ml water
  5 ml 10x TBE
  7.5 ml glycerol
  + 714 祃 10% APS + 17.2 祃 TEMED
References:
Mullis, K. and F. A. Faloona. (1987). "Specific synthesis of DNA in vitro via a polymerase catalyzed chain reaction."Meth. in Enzymol. 255:335-350.
Mullis, K, Faloona, F., Scharf, S., Saiki, R., Horn, G., and H. Erlich. (1986). "Specific enzymatic amplification of DNA in vitro: The polymerase chain reaction." Cold Spring Harbor Symposia on Quantitative Biology, Volume 51, Cold Spring Harbor Laboratory. p. 263-272.
Williams, J. F. (1989). "Optimization strategies for the polymerase chain reaction." Biotechniques 7(7):762-768.
PCR Technology: Principles and Applications for DNA Amplification. (1989). Erlich, H.A. (ed.), Stockton Press,

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cuturl('http://www.hgmp.mrc.ac.uk/GenomeWeb/nuc-primer.html')
cuturl('http://www.embl-heidelberg.de/ExternalInfo/geerlof/draft_frames/flowchart/clo_pcr_strategy/primer_design.html')
cuturl('http://bioweb.uwlax.edu/GenWeb/Molecular/Seq_Anal/Primer_Design/primer_design.htm')

PCR PRIMER DESIGN AND REACTION OPTIMISATION
Ed Rybicki, Department of Molecular and Cell Biology, University of Cape Town
copyright, 1992, 1996, 2001

Contents
·  Factors Affecting the PCR
o  Denaturing Temperature and Time
o  Annealing Temperature and Primer Design
o  Primer Length
o  Degenerate Primers
o  Elongation Temperature and Time
o  Reaction Buffer
o  Cycle Number
§  Nested Primer PCR
·  Labelling of PCR products with digoxygenin-11-dUTP
·  Helix Destabilisers / Additives
·  Useful Universal cDNA PCR Primer
·  A simple set of rules for primer sequence design
·  REFERENCES
Factors Affecting the PCR:
Denaturing Temperature and time
The specific complementary association due to hydrogen bonding of single-stranded nucleic acids is referred to as "annealing": two complementary sequences will form hydrogen bonds between their complementary bases (G to C, and A to T or U) and form a stable double-stranded, anti-parallel "hybrid" molecule. One may make nucleic acid (NA) single-stranded for the purpose of annealing - if it is not single-stranded already, like most RNA viruses - by heating it to a point above the "melting temperature" of the double- or partially-double-stranded form, and then flash-cooling it: this ensures the "denatured" or separated strands do not re-anneal. Additionally, if the NA is heated in buffers of ionic strength lower than 150mM NaCl, the melting temperature is generally less than 100oC - which is why PCR works with denaturing temperatures of 91-97oC.
A more detailed treatment of annealing / hybridisation is given in an accompanying page, together with explanations of calculations of complexity, conditions for annealing / hybridisation, etc.
Taq polymerase is given as having a half-life of 30 min at 95oC, which is partly why one should not do more than about 30 amplification cycles: however, it is possible to reduce the denaturation temperature after about 10 rounds of amplification, as the mean length of target DNA is decreased: for templates of 300bp or less, denaturation temperature may be reduced to as low as 88oC for 50% (G+C) templates (Yap and McGee, 1991), which means one may do as many as 40 cycles without much decrease in enzyme efficiency.
"Time at temperature" is the main reason for denaturation / loss of activity of Taq: thus, if one reduces this, one will increase the number of cycles that are possible, whether the temperature is reduced or not. Normally the denaturation time is 1 min at 94oC: it is possible, for short template sequences, to reduce this to 30 sec or less. Increase in denaturation temperature and decrease in time may also work: Innis and Gelfand (1990) recommend 96oC for 15 sec.

Annealing Temperature and Primer Design
Primer length and sequence are of critical importance in designing the parameters of a successful amplification: the melting temperature of a NA duplex increases both with its length, and with increasing (G+C) content: a simple formula for calculation of the Tm is
Tm = 4(G + C) + 2(A + T)oC.
Thus, the annealing temperature chosen for a PCR depends directly on length and composition of the primer. One should aim at using an annealing temperature (Ta) about 5oC below the lowest Tm of ther pair of primers to be used (Innis and Gelfand, 1990). A more rigorous treatment of Ta is given by Rychlik et al. (1990): they maintain that if the Ta is increased by 1oC every other cycle, specificity of amplification and yield of products <1kb in length are both increased. One consequence of having too low a Ta is that one or both primers will anneal to sequences other than the true target, as internal single-base mismatches or partial annealing may be tolerated: this is fine if one wishes to amplify similar or related targets; however, it can lead to "non-specific" amplification and consequent reduction in yield of the desired product, if the 3'-most base is paired with a target.
A consequence of too high a Ta is that too little product will be made, as the likelihood of primer annealing is reduced; another and important consideration is that a pair of primers with very different Tas may never give appreciable yields of a unique product, and may also result in inadvertent "asymmetric" or single-strand amplification of the most efficiently primed product strand.
Annealing does not take long: most primers will anneal efficiently in 30 sec or less, unless the Ta is too close to the Tm, or unless they are unusually long.
An illustration of the effect of annealing temperature on the specificity and on the yield of amplification of Human papillomavirus type 16 (HPV-16) is given below (Williamson and Rybicki, 1991: J Med Virol 33: 165-171).

Plasmid and biopsy sample DNA templates were amplified at different annealing temperatures as shown: note that while plasmid is amplified from 37 to 55oC, HPV DNA is only specifically amplified at 50oC.

Primer Length
The optimum length of a primer depends upon its (A+T) content, and the Tm of its partner if one runs the risk of having problems such as described above. Apart from the Tm, a prime consideration is that the primers should be complex enough so that the likelihood of annealing to sequences other than the chosen target is very low. (See hybridn.doc).
For example, there is a ¼ chance (4-1) of finding an A, G, C or T in any given DNA sequence; there is a 1/16 chance (4-2) of finding any dinucleotide sequence (eg. AG); a 1/256 chance of finding a given 4-base sequence. Thus, a sixteen base sequence will statistically be present only once in every 416 bases (=4 294 967 296, or 4 billion): this is about the size of the human or maize genome, and 1000x greater than the genome size of E. coli. Thus, the association of a greater-than-17-base oligonucleotide with its target sequence is an extremely sequence-specific process, far more so than the specificity of monoclonal antibodies in binding to specific antigenic determinants. Consequently, 17-mer or longer primers are routinely used for amplification from genomic DNA of animals and plants. Too long a primer length may mean that even high annealing temperatures are not enough to prevent mismatch pairing and non-specific priming.
Degenerate Primers
For amplification of cognate sequences from different organisms, or for "evolutionary PCR", one may increase the chances of getting product by designing "degenerate" primers: these would in fact be a set of primers which have a number of options at several positions in the sequence so as to allow annealing to and amplification of a variety of related sequences. For example, Compton (1990) describes using 14-mer primer sets with 4 and 5 degeneracies as forward and reverse primers, respectively, for the amplification of glycoprotein B (g from related herpesviruses. The reverse primer sequence was as follows:
TCGAATTCNCCYAAYTGNCCNT
where Y = T + C, and N = A + G + C + T, and the 8-base 5'-terminal extension comprises a EcoRI site (underlined) and flanking spacer to ensure the restriction enzyme can cut the product (the New England Biolabs catalogue gives a good list of which enzymes require how long a flanking sequence in order to cut stub ends). Degeneracies obviously reduce the specificity of the primer, meaning mismatch opportunities are greater, and background noise increases; also, increased degeneracy means concentration of the individual primers decreases; thus, greater than 512-fold degeneracy should be avoided. However, I have used primers with as high as 256- and 1024-fold degeneracy for the successful amplification and subsequent direct sequencing of a wide range of Mastreviruses against a background of maize genomic DNA (Rybicki and Hughes, 1990).


Primer sequences were derived from multiple sequence alignments; the mismatch positions were used as 4-base degeneracies for the primers (shown as stars; 5 in F and 4 in R), as shown above. Despite their degeneracy, the primers could be used to amplify a 250 bp sequence from viruses differing in sequence by as much as 50% over the target sequence, and 60% overall. They could also be used to very sensitively detect the presence of Maize streak virus DNA against a background of maize genomic DNA, at dilutions as low as 1/109 infected sap / healthy sap (see below).

Some groups use deoxyinosine (dI) at degenerate positions rather than use mixed oligos: this base-pairs with any other base, effectively giving a four-fold degeneracy at any postion in the oligo where it is present. This lessens problems to do with depletion of specific single oligos in a highly degenerate mixture, but may result in too high a degeneracy where there are 4 or more dIs in an oligo.