The products of a PCR reaction - especially when this is done on eukaryotic genomic DNA, and when using degenerate primers - often contain a mixture of discrete-sized bands, one of which is the "right" one, while the others represent products of "non-specific" priming. It can be a problem to obtain the correct band in any state approaching purity while maintaining yield, and attempting to purify the band by cloning all the reaction products and then probing the library for the correct DNA can be extraordinarily tedious.
I have applied a simple "core sampling" procedure - involving "coring" an agarose sample out of a gel, and using it as template for another round of PCR - to get around this problem, and obtain unique bands from initially messy backgrounds. Of course, having a visible band of the size expected does help; however, the technique may be used on faith on "right-sized" invisible bands if need be.
1. Run products of a PCR amplification on 1-2% TBE agarose gel, as two or more replicate lanes.
2. Cut off 1 lane - flanked by marker DNA if desired, and notched to allow re-orientation with remainder of gel - and stain in preferred ethidium bromide concentration (I use 50 ng/ml for 10 min).
3. View excised stained piece on 254nm UV box for maximum senssitivity; notch or stab correct band(s) in sample lane.
4. Prepare "core samplers": using gloves and sterile scissors and cut off about 5mm from the tip of as many sterile yellow pipette tips (we use Gilson tips) as you will need for samples.
5. Align stained marked segment with remainder of gel. Use "core samplers" to stab out one or more cores of agarose from the centre of bands of interest, using stabbed/notched gel lane as reference: a standard gel should give about 10ul per core.
6. Stain remainder of gel, view and photograph at 254nm to ensure correct regions were sampled.
NOTE: IT IS POSSIBLE TO QUICKLY CORE A STAINED GEL DIRECTLY ON A 305 OR EVEN A 254 NM UV BOX; HOWEVER, MORE THAN A FEW SECONDS OF EXPOSURE RESULTS IN CROSS-LINKING AND NO AMPLIFICATION
7. Use core samples as substrate in PCR reactions: I make up 40ul/reaction of reaction mix, and allow 10ul per core. Simply add core to mix, vortex a little, spin down, cover with mineral oil. PCR according to taste (not inhibited by presence of a little bromophenol blue or of 50ng/ml ethidium bromide).
8. At end of PCR: if you allow the tubes to cool down the reaction mix will set: 2%-odd agarose diluted 1/5 sets quite well! This is no problem for gel running as you then end the PCR on a 10 min 72oC cycle, and load the sample into wells of a gel BEFORE submerging the gel: sample will set in the wells and not float out.
9. If you wish to extract DNA, end at 72oC and add 50ul pre-warmed phenol / 8-OH-quinoline and vortex, add 100ul chloroform / isoamyl alcohol (24:1), vortex, spin: agarose should be in the phenol/CHCl3 phase. ALTERNATIVELY: take off mineral oil using 50ul CHCl3, take out plug of solidified sample and wash in TE, then put into 0.5ml Eppendorf-type vial with some siliconised glass wool at bottom, and a small needle hole. Put little Eppi in big Eppi without a lid, and spin 6000 rpm 10 min (a la Heery et al., 1990; TIG 6(6):173). Collect filtrate, clean up by phenol/CHCl3 and isopropanol/ammonium acetate ppte (1 vol IP, 0.2 vol of 10M ammonium acetate).
I have successfully re-amplified a unique 500bp band from a background of many bands up to 1.5kb from a cDNA PCR of cauliflower mosaic virus 35S RNA in total turnip RNA extract, and a 150bp band from a background of bands going up to 3kb from an amplification of Arabidopsis total genomic DNA using thoroughly degenerate primers - in the latter case, to a point where it could be sequenced directly (using same primers) after a subsequent amplification after purification from a gel plug as above.
The method has advantages over a previously-described toothpicking procedure in that a core sample is generally of defined volume, may be stored indefinitely, and provides material for multiple re-amplifications.
0.5 ug specific primer [eg. 0.3ul of 288uM=1.9ug/ul 20-mer]
1/20 vol DMSO [dimethyl sulphoxide]
DEPC-H20 to 22ul/rxn:
REMEMBER TO ALLOW FOR 10ul/rxn FOR SAMPLE!!
Mix everything together, leave on ice
HEAT SAMPLE MIX AT 65 C FOR 3 MIN, --> WET ICE
ADD 10ul RTase MIX / SAMPLE
MIX BY VORTEXING, HEAT SAMPLES AT 42 C 60 min / 52 C 30 min
Polymerase Chain Reaction (modified from PCR Protocols)
Reaction Mix:
It is again convenient to make up a master mix to be aliquotted out to amplify all samples, if they are to use the same primers. If not, modify master mix by simply leaving out primers.
Total volume: 50ul / rxn x N + 5-10% for wastage
Reaction master mix:
1/10 vol Cetus/ Promega /other 1 0x buffer
1/50 - 1/25 vol 2.5mM stock dNTPs [--> 50-100uM]
1/20 vol 10uM forward (=RNA sense) primer [--> 0.5uM]
[NB: no reverse primer needed IF THIS IS THE.SAME AS cDNA PRIMER as
residual cDNA synthesis primer concn. is +/- 5uM]
OPTIONAL: 1/20 - 1/10 vol DMSO
0.5ul of 5U/ul Taq polymerase / 100ul rxn mix
REMEMBER TO ALLOW FOR 5ul/rxn OF SAMPLE!!
MIX MASTER RXN MIX, LEAVE ON ICE.
Aliquot out 45ul / PCR rxn vial
Add 5ul sample / vial from reverse transcription reaction mix
Add 50ul / vial mineral oil (NB: new upipette tips each 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 / hybridization, 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.
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(s). 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> 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.
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.
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 (gB) 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(s), 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.
This is normally 70 - 72oC, for 0.5 - 3 min. Taq actually has a specific activity at 37oC which is very close to that of the Klenow fragment of E coli DNA polymerase I, which accounts for the apparent paradox which results when one tries to understand how primers which anneal at an optimum temperature can then be elongated at a considerably higher temperature - the answer is that elongation occurs from the moment of annealing, even if this is transient, which results in considerably greater stability. At around 70oC the activity is optimal, and primer extension occurs at up to 100 bases/sec. About 1 min is sufficient for reliable amplification of 2kb sequences (Innis and Gelfand, 1990). Longer products require longer times: 3 min is a good bet for 3kb and longer products. Longer times may also be helpful in later cycles when product concentration exceeds enzyme concentration (>1nM), and when dNTP and / or primer depletion may become limiting.
and/or non-ionic detergents such as Tween-20 or Nonidet P-40 or Triton X-100 (0.05 - 0.10% v/v)
(Innis and Gelfand, 1990). Modern formulations may differ considerably, however - they are also generally proprietary.
PCR is supposed to work well in reverse transcriptase buffer, and vice-versa, meaning 1-tube protocols (with cDNA synthesis and subsequent PCR) are possible (Krawetz et al., 19xx; Fuqua et al., 1990).
Higher than 50mM KCl or NaCl inhibits Taq, but some is necessary to facilitate primer annealing.
[Mg2+] affects primer annealing; Tm of template, product and primer-template associations; product specificity; enzyme activity and fidelity. Taq requires free Mg2+, so allowances should be made for dNTPs, primers and template, all of which chelate and sequester the cation; of these, dNTPs are the most concentrated, so [Mg2+] should be 0.5 - 2.5mM greater than [dNTP]. A titration should be performed with varying [Mg2+] with all new template-primer combinations, as these can differ markedly in their requirements, even under the same conditions of concentrations and cycling times/temperatures.
Some enzymes do not need added protein, others are dependent on it. Some enzymes work markedly better in the presence of detergent, probably because it prevents the natural tendency of the enzyme to aggregate.
Primer concentrations should not go above 1uM unless there is a high degree of degeneracy; 0.2uM is sufficient for homologous primers.
Nucleotide concentration need not be above 50uM each: long products may require more, however.
The number of amplification cycles necessary to produce a band visible on a gel depends largely on the starting concentration of the target DNA: Innis and Gelfand (1990) recommend from 40 - 45 cycles to amplify 50 target molecules, and 25 - 30 to amplify 3x105 molecules to the same concentration. This non-proportionality is due to a so-called plateau effect, which is the attenuation in the exponential rate of product accumulation in late stages of a PCR, when product reaches 0.3 - 1.0 nM. This may be caused by degradation of reactants (dNTPs, enzyme); reactant depletion (primers, dNTPs - former a problem with short products, latter for long products); end-product inhibition (pyrophosphate formation); competition for reactants by non-specific products; competition for primer binding by re-annealing of concentrated (10nM) product (Innis and Gelfand, 1990).
If desired product is not made in 30 cycles, take a small sample (1ul) of the amplified mix and re-amplify 20-30x in a new reaction mix rather than extending the run to more cycles: in some cases where template concentration is limiting, this can give good product where extension of cycling to 40x or more does not.
A variant of this is nested primer PCR: PCR amplification is performed with one set of primers, then some product is taken - with or without removal of reagents - for re-amplification with an internally-situated, "nested" set of primers. This process adds another level of specificity, meaning that all products non-specifically amplified in the first round will not be amplified in the second. This is illustrated below:
This gel photo shows the effect of nested PCR amplification on the detectability of Chicken anaemia virus (CAV) DNA in a dilution series: the PCR1 just detects 1000 template molecules; PCR2 amplifies 1 template molecule (Soiné C, Watson SK, Rybicki EP, Lucio B, Nordgren RM, Parrish CR, Schat KA (1993) Avian Dis 37: 467-476).
Labelling of PCR products with digoxygenin-11-dUTP
(DIG; Roche) need be done only in 50uM each dNTP, with the dTTP substituted to 35% with DIG-11-dUTP. NOTE: that the product will have a higher MW than the native product! This results in a very well labelled probe which can be extensively re-used, for periods up to 3 years. See also here.
With NAs of high (G+C) content, it may be necessary to use harsher denaturation conditions. For example, one may incorporate up to 10% (w or v/v) :
dimethyl sulphoxide (DMSO),
dimethyl formamide (DMF),
urea
or formamide
in the reaction mix: these additives are presumed to lower the Tm of the target NA, although DMSO at 10% and higher is known to decrease the activity of Taq by up to 50% (Innis and Gelfand, 1990; Gelfand and White, 1990).
Additives may also be necessary in the amplification of long target sequences:DMSO often helps in amplifying products of >1kb. Formamide can apparently dramatically improve the specificity of PCR (Sarkar et al., 1990), while glycerol improves the amplification of high (G+C) templates (Smith et al., 1990).
Polyethylene glycol (PEG) may be a useful additive when DNA template concentration is very low: it promotes macromolecular association by solvent exclusion, meaning the pol can find the DNA.
cDNA PCR
A very useful primer for cDNA synthesis and cDNA PCR comes from a sequencing strategy described by Thweatt et al. (1990): this utilised a mixture of three 21-mer primers consisting of 20 T residues with 3'-terminal A, G or C, respectively, to sequence inside the poly(A) region of cDNA clones of mRNA from eukaryotic origin. I have used it to amplify discrete bands from a variety of poly(A)+ virus RNAs, with only a single specific degenerate primer upstream: the T-primer may anneal anywhere in the poly(A) region, but only molecules which anneal at the beginning of the poly(A) tail, and whose 3'-most base is complementary to the base next to the beginning of the tail, will be extended.
eg: 5'-TTTTTTTTTTTTTTTTTTTTTTTTT(A,G,C)-3'
works for amplification of Potyvirus RNA, and eukaryotic mRNA
3.primers should end (3') in a G or C, or CG or GC: this prevents "breathing" of ends and increases efficiency of priming;
4.Tms between 55-80oC are preferred;
5.runs of three or more Cs or Gs at the 3'-ends of primers may promote mispriming at G or C-rich sequences (because of stability of annealing), and should be avoided;
6.3'-ends of primers should not be complementary (ie. base pair), as otherwise primer dimers will be synthesised preferentially to any other product;
7.primer self-complementarity (ability to form 2o structures such as hairpins) should be avoided.
Examples of inter- and intra-primer complementarity which would result in problems:
Screen shots taken from analyses done using DNAMAN (Lynnon Biosoft, Quebec, Canada).
This is best done using ssDNA generated by asymmetric PCR, and the "limiting" primer for sequencing. However, efficient sequencing of dsDNA generated by normal PCR is possible using the modification to the SequenaseTM protocol published by Bachmann et al. (1990) (NAR 18: 1309). CLEAN DNA is resuspended in sequencing buffer containing 0.5% Nonidet P-40 and 0.5% Tween-20 and sequencing primer, denatured by heating to 95oC for 5 min, snap-cooled on wet ice, and sequenced by the "close-to-primer" protocol (eg: dilute extension mixes).
T-A Cloning Strategy: Taq and other polymerases seem to have a terminal transferase activity which results in the non-templated addition of a single nucleotide to the 3'-ends of PCR products. In the presence of all 4 dNTPs, dA is preferentially added; however, use of a single dNTP in a reaction mix results in (relatively inefficient) addition of that nucleotide. This complicates cloning, as the supposedly blunt-ended PCR product often is not, and blunt-ended cloning protocols often do not work or are very inefficient. This can be remedied by incubation of PCR products with T4 DNA pol or Klenow pol, which "polishes" the ends due to a 3'->5' exonuclease activity (Lui and Schwartz, 1992; BioTechniques, 20: 28-30). However, this terminal transferase activity is also the basis of a clever cloning strategy: this uses Taq pol to add a single dT to the 3'-ends of a blunt-cut cloning vector such as pUC or pBluescriptTM, and simple ligation of the PCR product into the now "sticky-ended" plasmid (Marchuk et al., 1990; NAR 19: 1156).
Incorporation of Restriction Sites in Primers:
Although this may be rendered simple by incorporating the same or different restriction sites at the 5'-ends of PCR primers - which allows generation of sticky ends and straightforward cloning into appropriate vectors - these should have AT LEAST two additional bases 5' to the recognition sequence to ensure that the enzymes will in fact recognise the sequence - and it is often found that even when this is done, the efficiency of cutting of fresh product is next to zero. This can sometimes be remedied by incubating fresh product with Proteinase K (to digest off tightly-attached Taq pol), but often is not. A solution to the problem is to use the "Klenow-Kinase-Ligase" (KKL) method: this involves "polishing" products with Klenow, kinasing them to get 5'-phosphorylation (NB: OLIGONUCLEOTIDE PRIMERS NORMALLY HAVE NO 5'-PHOSPHATES!!!), ligating the fragments together to get concatemers, then restricting these with the appropriate restriction enzymes to generate the sticky-ended fragments suitable for cloning (Lorens, 1991; PCR Methods and Applications, 1: 140-141).
• Getting rid of mineral oil: simply add 50ul of chloroform to the reaction vial, vortex and centrifuge briefly, and remove the "hanging droplet" of AQUEOUS solution with a micopipette.
• Getting rid of wax or Vaseline: simply "spear" wax gem and remove; do as for oil or bottom-puncture tube and blow out aqueous drop for Vaseline.
• Cleaning-up DNA: 3 options
a protocol which gives DNA that is clean enough for sequencing is the following: increase volume to 100ul with water, add 10M ammonium acetate soln. to 0.2M final concentration (1/5th volume), add equal volume of isopropanol (propan-2-ol), leave on bench 5 min, centrifuge 20 min at 15 000 rpm, remove liquid using pipette, resuspend in 100ul water or TE, repeat precipitation.
Simply do a phenol-CHCl3 extraction (add 20ul phenol to aqueous phase, vortex, add 50ul CHCl3, vortex, centrifuge, remove UPPER aqueous phase, repeat CHCl3 extraction).
Make aqueous phase up to 400ul, and spin through Millipore Ultrafree-MC NMWL 30 000 cartridges (at 6000 rpm in microcentrifuge), wash through with 2x400ul water, collect +/-20ul sample: this is pure enough for sequencing.
NOTE:
Product is clean enough for restriction digest immediately after reaction; however, phenol-chloroform clean-up is recommended as a minimum.
PCR products may be very conveniently labelled with digoxigenin-11-dUTP (Boehringer-Mannheim) by incorporating the reagent to 10-35% final effective dTTP concentration in a nucleotide mix of final concentration 50-100uM dNTPs (Emanual, 1991; Nucleic Acids Res 19: 2790). This allows substitution to a known extent of probes of exactly defined length, which in turn allows exactly defined bybridisation conditions. It is also the most effective means of labelling PCR products, as it is potentially unsafe and VERY expensive to attempt to do similarly with 32P-dNTPs, and nick-translation or random primed label incorporation are unsuitable because the templates are often too small for efficient labelling.
Make a DIG-dNTP mix for PCR as follows:
DIG NUCLEOTIDE MIX CONCENTRATIONS
Dig-11-dUTP 350 uM
dTTP 650 uM
dATP 1 mM
dCTP 1 mM
dGTP 1 mM
For each 50 ul of probe synthesized, a 1/10 dilution is made of the DIG-nucleotide mix when added to the other reagents as described above. The products may be analyzed by agarose gel electrophoresis - NOTE: PRODUCTS ARE LARGER THAN NON-SUBSTITUTED PRODUCT - and detected directly on blots immunologically. Probes can be used as 5-10 ul aliquots directly from PCR product mixes, mixed with hybridisation mix and denatured. Probes can be re-used up to 10 times, stored frozen in between experiments and boiled to denature.
Take 1/10th - 1/3rd of the reaction mix CAREFULLY from under the oil or from under the Vaseline or solidified wax, using a micropipette with plugged tip, IN AN AREA AWAY FROM YOUR PCR PREPARATION AREA!
Mix this with some gel loading buffer(1:1 - 1:5 mix:loading buffer): this is TBE containing 10 - 20% glycerol or sucrose and a dash of bromophenol blue (BPB) tracking dye.
Load 5 - 30ul of sample into wells of 0.8 - 3.0% submarine agarose gel made up in TBE, preferably containing 50ng/ml ethidium bromide.
Run at 80 -120 volts (not too slow or small products diffuse; not too fast or bands smear) until BPB reaches end of gel (large products) or 2/3 down gel (small products). Use DNA markers going from 2kb down to 100 bp or less (recommend BM PCR markers). View on UV light box at 254 - 300 nm, photo 1 - 5 sec.
NOTE:
Small products are best seen on 3% agarose gels that have been run fast (eg: 100 volts), with BPB run to ½ - 2/3 down the gel. It is best to include EthBr in the gel AND in the gel buffer, as post-electrophoresis staining can result in band smearing due to diffusion, and if there is no EthBr in the buffer the dye runs backwards out of the gel, and smaller bands are stripped of dye and are not visible.
NUSIEVE TM gel (FMC Corp) can also be used for small products - better resolution than agarose.
Polyacrylamide gels can be silver stained by simple protocols for extreme sensitivity of detection.
Gels can be blotted directly after soaking in 0.5M NaOH / 1.5M NaCl for 10-20 min: "dry blotting" works well (eg: gel is over- and under-layered with paper towel stacks and pressed; bands transfer up and down), as does classic "Southern" blotting. Bands blotted in this way are already covalently fixed onto nylon membranes, and simply need a rinse in 5xSSPE before prehybridisation.
The example shown is of detection of Human papillomavirus type 16 (HPV-16) DNA amplified from cervical biopsy samples (Williamson A-L, Rybicki EP (1991) Detection of genital human papillomaviruses by polymerase chain reaction amplification with degenerate nested primers. J Med Virol 33: 165-171). The left panel is a photo of an EthBR-stained 2% agarose gel; the right is an autoradiograph of a Southern blot probed with 32P-labelled HPV-16 DNA. Note how much more sensitive blotting is, and how much more specific the detection is.
i) Oligonucleotide primers are generally supplied as "so many OD units/ml" - but what does this mean, in terms of mg/ml, or mmol/ml, etc?
Given: a primer is Y nucleotides (nt) long;
Given: the MW of ssDNA is (330 daltons per nt) x (length in nt) (Sambrook et al., 1989; p. C.1);
Given: the concentration of primer (=ssDNA) producing an OD of 1 at 254 nm in a 1 cm cuvette, is 37 ug/ml;
Then: the MW of the primer is 330.Y daltons
And: X OD/ml = 37.X ug/ml = 37.X mg/l = 37.X /330.Y mM = 37.X.1000/330.Y uM
For example:
B 88/77 primer - a 17-mer oligodeoxynucleotide - as supplied is 12.6 OD units/ml. We need to make a 5 uM stock solution for PCR.
MW: 17 x 330 = 5610
Concentration: 12.6 OD x 37 ug/ml = 466 ug/ml = 466 mg/l = 0.466 g/l
Molarity: 0.466/5610 = 0.000083 Molar = 83 uM
Therefore: we need 5 ul of oligo stock solution in 83 ul (+78 ul water) to make a 5 uM solution (if 1 ul in 83 ul gives a 1 uM soln...)
ii) Calculation of amounts for PCR reactions: if we need a final concentration of 0.5 uM oligo in the PCR reaction mix (final volume 50 ul), we add 5 ul of 5 uM stock to the reaction mix (1/10 final dilution).
Stocks of nucleotides for PCR (or other procedure) are NEARLY ALWAYS dNTPs (deoxynucleotides), and concentrations is almost always given in EACH dNTP: that is, the given concentration is EACH nucleotide in the mix, NOT the total concentration. This means that a 2.5 mM dNTP mix for PCR contains 2.5 mM of EACH dNTP, and 10 mM TOTAL dNTPs.
Example:
i) Make up a 2.5 mM stock solution of dNTPs from stock 100 mM individual dNTPs, supplied by Promega:
FIRST mix equal volumes of each nucleotide (eg: 50 ul): this gives you 200 ul of 25 mM mixed dNTPs (Remember: concn. expressed in EACH dNTP).
THEN dilute this (or aliquot) 1/10 with WATER - aliquot into 100 ul amounts and freeze.
ii) Prepare a 1 mM stock of dNTPs with dTTP substituted to 10% (w/w) by digoxigenin-11-dUTP (DIG-dUTP) for use as a labelling mix for PCR labelling of PCR products:
GIVEN:
DIG-dUTP supplied (by Boehringer Mannheim) at 25 nmol/25ul = 1 umol/ml = 1mM; final concentration of DIG-dUTP must be 1/10th that of other nucleotides, and [DIG-dUTP] + [dTTP] must = [any other dNTP]. Therefore to get a 1 mM dNTP stock one must dilute DIG-dUTP stock 1/10.
FIRST dilute separate 100 mM dNTP stocks to 10 mM (eg. 5 ul to 50 ul, in water).
THEN mix equal volumes (eg. 10 ul) of 10 mM dCTP, dGTP and dATP stock, and 9/10ths volume of dTTP (9 ul). Add equal volume (eg. 10 ul) of of 1 mM DIG-dUTP.
THEN add water to 10 vol (=100 ul; add 51 ul): final concentration each dNTP = 1 mM; final concn DIG-dUTP = 0.1 mM, and of dTTP = 0.9 mM.
iii) USE mix made above at 50 uM each dNTP in a PCR reaction mix, final volume 25ul:
NEED to dilute mix 1/20; therefore use 1.25 ul dNTP labelling mix per 25 ul reaction volume (1/20 = 5/100 = 1.25/25).
To make mastermix: multiply amount of dNTP per reaction by number of reactions.
In certain circumstances one wishes to avoid mixing primers and target DNA at low temperatures in the presence of Taq polymerase: Taq pol is almost as efficient as Klenow pol at 37oC; consequently, if primers mis-anneal at low temperature prior to initial template denaturation, "non-specific" amplification may occur. This may be avoided by only adding enzyme after the initial denaturation, before the reaction cools to the chosen annealing temperature. This is most conveniently done by putting wax "gems"TMinto the reaction tube after addition of everything except enzyme, then putting enzyme on top of the gem: the wax melts when the temperature reaches +/-80oC, and the enzyme mixes with the rest of the reaction mix while the molten wax floats on top and seals the mix, taking the place of mineral oil. Information is that "gems" may be substituted by VaselineTM.
• Primers: 0.2 - 1.0 uM • Nucleotides: 50 - 200 uM EACH dNTP • Dimethyl sulphoxide (DMSO): 0 - 10% (v/v) • Taq polymerase: 0.5 - 1.0 Units/50ul rxn Target DNA: 1 ng - 1 ug (NB: higher concn for total genomic DNA; lower for plasmid / purified DNA / virus DNA target)
Buffer: use proprietary or home-made 10x rxn mix; eg: Cetus, Promega. This should contain: minimum of 1.5mM Mg2+, usually some detergent, perhaps some gelatin or BSA. Promega now supply 25mM MgCl2, to allow user-specified [Mg2+] for reaction optimisation with different combinations of primers and targets.
MAKE POOLED MASTER MIX OF REAGENTS IN ABSENCE OF DNA using DNA-free pipette, then dispense to individual tubes (using DNA-free pipette), and add DNA to individual reactions USING PLUGGED TIPS.
OVERLAY REACTIONS WITH 50UL OF HIGH-QUALITY LIQUID PARAFFIN OR MINERAL OIL to ensure no evaporation occurs: this changes reactant concentrations. NOTE: latest wisdom has it one can use VASELINE - this also allows "HOT START" PCR.
NOTE
USE PLUGGED PIPETTE TIPS: prevents aerosol contamination of pipettes. Use of detergents is recommended only for Taq from Promega (up to 0.1% v/v, Triton X-100 or Tween-20). DMSO apparently allows better denaturation of longer target sequences (>1kb) and more product.
DO NOT USE SAME PIPETTE FOR DISPENSING NUCLEIC ACIDS AS YOU USE FOR DISPENSING REAGENTS
Remember sample volume should not exceed 1/10th reaction volume, and sample DNA/NTP/primer concentrations should not be too high as otherwise all available Mg2+ is chelated out of solution and enzyme reactivity is adversely affected. Any increase in dNTPs over 200uM means [Mg2+] should be re-optimised.
AVOID USING EDTA-CONTAINING BUFFERS AS EDTA CHELATES Mg2+ Low primer, target, Taq, and nucleotide concentrations are to be favoured as these generally ensure cleaner product and lower background, perhaps at the cost of detection sensitivity. Recommended Reaction Conditions:
Initial Conditions:
Initial denaturation at start: 92 - 97oC for 3 - 5 min. If you denature at 97oC, denature sample only; add rest of mix after reaction colls to annealing temperature (prevents premature denaturation of enzyme). Initial annealing temperature: as high as feasible for 3 min (eg: 50 - 75oC). Stringent initial conditions mean less non-specific product, especially when amplifying from eukaryotic genomic
DNA.
Initial elongation temperature: 72oC for 3 - 5 min. This allows complete elongation of product on rare templates. (see also here) Temperature Cycling: • 92 - 94oC for 30 - 60 sec (denature) • 37 - 72oC for 30 - 60 sec (anneal) • 72oC for 30 - 60 sec (elongate) (60 sec per kb target sequence length) • 25 - 35 cycles only (otherwise enzyme decay causes artifacts) • 72oC for 5 min at end to allow complete elongation of all product DNA
NOTE:
"Quickie" PCR is quite feasible: eg, [94oC 30 sec / 45oC 30 sec / 72oC 30 sec] x 30, for short products (200 - 500 bp). YOU CAN USE GLYCEROL IN THERMAL CYCLER REACTION TUBE HOLES TO ENSURE GOOD THERMAL CONTACTS DON'T RUN TOO MANY CYCLES: if you don't see a band with 30 cycles you probably won't after 40; rather take an aliquot from the reaction mix and re-PCR with fresh reagents
Introduction:Mycoplasma synoviae (MS) is a major pathogen of chickens and turkeys causing upper respiratory infection and synovitis in chickens and turkeys. Additionally, MS infection in turkeys has been associated with increased condemnation at processing due to airsacculitis, as well as with breast blisters and green discoloration of liver, which results in slower processing and further economic loss. Because of the high value of individual breeder hens and because of their ability to infect progeny by vertical transmission, economic losses are potentially most severe when MS infections occur in breeder flocks. Therefore a reasonable strategy for control of MS is to establish MS-free breeder flocks. Diagnosis of MS infections is often carried out by serological procedures but non specific reactions or insensitivity of the serological detection of MS infection often limits the effectiveness of these procedures. Isolation of the organism is also used but may be unsuccessful due to bacterial contamination, mixed mycoplasma infections or other reasons. Successful isolation may require 1-3 weeks. Species specific PCR tests are used based on 16s rRNA and vlhA genes. The major advantages of Real-Time PCR over standard PCR assay are the one tube and one time handling with reading of the reaction during the assay and less time needed to perform the test.
Objective:To develop and optimize real time PCR assay for quantitative detection of Mycoplasma synoviae infections from clinical samples.
Methodology:M. synoviae strains/isolates from chicken and turkey will be procured for standardization of the real time PCR assay. Clinical samples (trachea, air sac, synovial fluid) from affected chicken/turkey will be collected. Nucleic acids will be extracted directly from different M. synoviae strains/isolates and clinical samples and will be subjected to real-time PCR. Primer and fluorescent tagged probe set used in this investigation will be developed and characterized. The amplification of DNA fragment from the suspect samples detected by monitoring the increase in fluorescence from the dye labeled MS-specific probe will be investigated
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