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Andrew Krohn's PCR troubleshooting page
Ph.D student, Northern Arizona University Department of Biological Sciences
Disclaimer: I am by no means an expert at PCR. I learn new things all the time. Everything covered here is a mixture of experiences I have gained (particularly in the Plumley, Winton, Riday and Gehring labs). The excellent three volume set, Molecular Cloning by Sambrook & Russell and Octavian Henegariu's wonderful PCR site have been tremendously helpful to me at times. As well, I consider Andrew Lang an inspiration in lab technique which need not be hindered by contemporary laboratory superstitions.
What is PCR?
Polymerase chain reaction or PCR is an improved method for amplifying specific DNA sequences. It was modified to its present form in the 1980s by Kary Mullis for which he was awarded the Nobel Prize in Chemistry in 1993. The process begins with a heating step (usually ~95 degrees C) that denatures double-stranded DNA into single-stranded components. The second step cools the reaction to an annealing temperature (sequence dependent, usually somewhere betwee 45 and 65 degrees C). The third step raises the temperature of the reaction to where Taq DNA polymerase is active (usually between 65 and 72 degrees C). The temperature is maintained at the third step long enough for the desired sequence to be copied completely and the three step process repeats. As the process repeats successively, the target sequence is amplified logarithmically. A typical PCR program will include between 25 and 40 cycles. Though reactions are never 100% efficient, this means that a tiny amount of DNA, say 5 ng (nanograms) can be used to produce several ug (micrograms) of DNA in just a few hours. Amplification of specific sequences allows researchers to study DNA in more detail and "read" the DNA bases (A, C, T, and G) found within their sequence of interest. This is useful for studying many aspects of biology such as human health, population genetics, ecology, and evolution to name just a few.
PCR ingredients:
In order to have continued success in performing PCR on different loci in different organisms or in slightly more advanced techniques such as AFLP or multiplex PCR, one must have a thorough understanding of the contents of their PCR reaction. In some cases the recipe must be tweaked slightly in order to get the desired results. The following table describes typical PCR reaction contents.
| Reaction component |
Concentration range |
Concentration I typically use |
Reason for inclusion |
| Tris-Cl |
|
10 mM |
Stabilizes pH. |
| KCl |
30-100 mM |
50 mM |
Concentration greater than 50 mM may increase the efficiency of your reaction. |
| MgCl2 |
1.0-4.0 mM |
2.0 mM |
Cofactor required for Taq polymerase activity. High concentrations may actually inhibit Taq DNA polymerase and decrease product specificity. |
| Gelatin |
|
0.0001% v/v |
Increases macromolecular crowding raising the rate of higher order kinetics (e.g. polymerase + DNA + MgCl2 + dNTPs). |
| dNTPs (ea) |
50-500 uM |
200 uM |
The constituent bases of DNA, therefore required for new synthesis. Higher concentrations may contribute to production of non-specific products and/or sequestration of available MgCl2. |
| Taq DNA polymerase |
0.005-0.05 U/uL |
0.015 U/ul |
The molecular machine itself! Lower concentrations reduce production of non-specific products. |
| Primers (ea) |
100-500 nM |
200 nM |
Synthetic oligonucleotides with sequences complementary to the sequence you wish to amplify. Lower concentrations reduce production of non-specific products |
| Template DNA |
~1-100 ng |
5-10 ng |
The DNA from which you are producing an amplified product. Higher concentrations are required as template complexity increases. Too high of concentration can result in production of non-specific products. |
| Total volume |
6-10 uL |
|
|
Other components sometimes used in PCR reactions are bovine serum albumin (BSA), dithiothreitol (DTT), sodium dodecyl sulfate (SDS), glycerol, sucrose, and dimethyl sulfoxide (DMSO). I probably missed an alternative ingredient or three.
Thermal cycle:
The temperatures you use in your thermal cycle can have a profound outcome on your PCR reaction.
| Step |
Common range |
Range I typically use |
Reason |
| Initial denature |
92-96C, 1-8 min |
95C, 2 min |
Denatures DNA. Sometimes an initial denature is necessary for amplification from complex DNA. |
| Denature |
92-96C, 10-60 sec |
95C, 30 sec |
Denatures DNA during cycle |
| Anneal |
42-65C, 30-90 sec |
50-62C, 30-90 sec |
Temperature at which your primers anneal to your chosen target. Lower temperature reduces annealing specificity. For complex targets or multiplex PCR, a longer annealing time may be necessary. |
| Extend |
65-72C, 45-90 sec |
65-68C, 45-90 sec |
Temperature at which Taq DNA polymerase is most active. Higher temperature corresponds with greater activity, but (anecdote here) a lower temperature can reduce errors made by the polymerase. |
| Final extension |
65-72C, 2-10 min |
68C, 5 min |
Gives the polymerase a chance to complete any unfinished strands of DNA in your reaction mixure. For many purposes this is probably unnecessary. Terminal adenines are also filled in where missing (Taq DNA polymerase always leaves a 3' A overhang). I do a lot of fragment analysis by capillary electrophoresis and use of the final extension helps reduce ambiguity in calling fragment sizes due (presumably) to incomplete terminal adenylation. |
| Cold hold |
4C, forever |
10C, forever |
Many people like to leave the thermal cycler at a cold temperature so when they leave a reaction on the cycler and go home at night, the reaction is the refrigerated until they arrive in the morning. This is another unnecessary step as the DNA should be quite stable in the PCR mixture at room temperature. I use an increased temperature as it is still cold enough to prevent small reaction volumes (6ul) from evaporating, but is less taxing on the thermal cycler. |
The above table represents the general model for PCR reactions and does not address 2-step protocols, temperature or heating ramps, temperature gradients, or other specialized cycling parameters.
Help! My PCR didn't work!
Every researcher has experienced failed PCR. This can happen for many reasons, but in my experience the reason most often responsible is poor quality DNA used as template. Most likely your DNA extraction didn't yield enough DNA or else it also carried through some cocktail of secondary metabolites from the organism you are studying which are inhibiting your reaction from proceeding. Sometimes template DNA is too concentrated. Take these steps to check your DNA quality:
- Use a spectrophotometer to read your DNA. Pay attention to the concentration. Is it high enough? I work with pines which have numerous large chromosomes and 5ng is sufficient for amplification of any locus in a 6uL reaction volume.
- Look at the 260/280 ratio. It should be in the 1.5-1.8 range. If not, you may have contaminating proteins in your DNA sample. A phenol/chloroform (sometimes just chloroform works fine) extraction should remedy this problem.
- Look at the 260/230 ratio. The higher the better. Low 260/230 ratio (less than 0.5 or so) may indicate contaminating secondary metabolites. This can be tricky to resolve, especially if you use kits for DNA extraction. I tend to use "homebrew" DNA extractions and I have found including polyvinyl pyrrolidone (MW 40,000) at 2% and polyethylene glycol (MW 8,000) at 10% helps remedy the secondary metabolite issue.
- Run some of your DNA sample on a gel so you can directly visualise it. Use a quantitative DNA ladder if you have it so you can estimate actual concentration (OD 260 is not always reliable). Is it at least 10,000kb (good)? Physical shearing (due to vortexing, pipeting etc) tends to degrade DNA in terms of size. Alternatively, contaminating DNases could do the same thing as can repeated freezing and thawing of your samples. Degraded DNA may yield a nice-looking spectrophotometer profile and yet no PCR is seemingly possible. If your DNA leaves streaks, you may have some protein or secondary metabolite contamination. Alternatively, maybe you just loaded too much sample on the gel.
- Try a "lazy-man's dilution series" PCR. Prepare master mix for a target you know works (maybe "universal" ribosomal primers). Plan to do 3 reactions per sample in 10uL volumes. Aliquot 9uL to each reaction well. To the first reaction, add 1uL template DNA. Mix well by inversion and spin down. Withdraw 1uL of the first reaction and add it to the second reaction as template (now 1/10 concentration of the first reaction). Mix well and spin down. To the third reaction, add 1 (1/100X) or 2 (1/50X) uL of the second reaction as template. Mix well, spin down, and run the reaction on the thermal cycler. Note: in case your DNA concentration is already low and by doing this you simply dilute PCR inhibitors to an acceptable level, make sure you run your PCR at least 35 cycles (I do 40).
- If the above doesn't work, consider your thermal cycle (sometimes it is wise to even consider this first). Is your annealing temperature low enough? It should be within 5 degrees C of the lower melting temperature (Tm) of your two primers. Is your extension time long enough? Conventional wisdom dictates that taq DNA polymerase synthesizes about 1000 bases per minute at 72 degrees C. I usually use a 68 degrees C extension temperature so I will fudge a few extra seconds to my extension times. Are you using enough cycles? I get useful product at even 25 cycles but if I need a lot of product or my DNA template is very dilute 35 or more cycles may be necessary.
- Degraded primers? Are your primers old? Have they been thawed and refrozen many many times? What are they dissolved in? Some people resuspend primers in water. This is OK for a while, but even purified laboratory water tends to be a little acidic. DNA is an acid and so is most stable in a slightly basic solution. For this reason, I always dissolve my primers in Tris-Cl pH 8.0 and my primers are always good for several years. I used to dissolve my primers in TE, but then I realized that by doing so I was effectively reducing the concentration of MgCl2 in my PCR reactions when I added the primers. The EDTA is there to inhibit any loose nucleases that somehow persist in your solutions. Since my technique is careful and sanitary, I decided to assume I was experiencing no random nuclease contamination. Several years later, everything is still great!
- Polymerase gone bad? If an absent-minded (drunk maybe?) grad student was performing PCR late one night, maybe they forgot to put stuff back in the freezer. I wouldn't know anything about that! Anyways, when all else fails, borrow some polymerase from someone else to eliminate this obnoxious possibility. Note: I always aliquot my polymerase master mix into small volumes (500 uL) so if this happens, not much is actually wasted.
PCR tips:
...coming soon
A reckless comparison of Taq DNA polymerases:
...coming soon
Questions? Comments? Contact me: alk224@nau.edu |