Methods and reagents: Wayward PCR primers

by Paul N. Hengen, Ph.D. *

Methods and reagents is a unique monthly column that highlights current discussions in the newsgroup bionet.molbio.methds-reagnts, available on the Internet. This month's column discusses the failure of PCR primers that have been kept a long time to amplify DNA. For details on how to partake in the newsgroup, see the accompanying box.

Some netters have noticed a loss in the ability to amplify DNA by the polymerase chain reaction (PCR) when using the same batch of primers over a period of time. PCR primers performed well shortly after being purchased, but after storage for several months as diluted stocks in water within a non-frost-free -20 degrees C freezer, these same primers failed to give the same amount of amplified product in identical reactions. If new primers were purchased, the reactions gave good results and acted just like the old primers did before storage.

Bad seed, bad storage or bad company?
The reason behind this phenomenon has been baffling netters as they search for a way to avoid buying a new set of primers every few months. One idea is that the failure occurs when doing long and accurate (LA) PCR, a sometimes difficult operation in itself, and that the carryover that contaminates products from earlier PCR samples used as template is to blame. This has been described by Wayne Barnes as the `bad seed' in an earlier Methods and reagents column [see TIBS 19 (1994), 341-342]. It could explain why PCR fails over time, but cannot explain why newly purchased primers would work.

Some netters became highly concerned about the storage conditions for their own primers when hearing of this, because larger projects requiring as many as several hundred sets of primers could cost enormous sums of money to replace.

One netter, Richard Maleszka (maleszka@rsbscentral.anu.edu.au), wrote about his experience with long-term storage of primers frozen at -25 degrees C in water at pH 7.0. He found that three-month-old primers of 22-24 nucleotides in length initially worked beautifully, but after a longer storage period they didn't yield any amplification product.

While others were calling for hard evidence that the primers are at fault, Mike Cooley (mbcooley@ucdavis.edu) wrote that he has been using nested PCR to detect very rare events, and is therefore pushing the sensitivity limit of PCR. He was unable to amplify fragments from DNA that had succeeded about a year previously under identical conditions. After ordering new oligos and performing side-by-side comparisons with both sets of oligos, he confirmed that the primers had `gone bad'.

Bad primers were also to blame for unreadable results in automated DNA sequencing reactions using the dye-terminator chemistry combined with cycle sequencing and subsequent analysis on the ABI model 373A sequencer. The same conditions had previously given good results with fresh primers.

Another netter observed loss of signal with 5'-fluorescent-tagged primers stored at a concentration of 10 pmol/ul in Tris/EDTA buffer for six months. Fluorescent signal strength was reduced three or more orders of magnitude depending on the primer set, compared with the same primers used under the identical conditions six months previously. However, it was not clear whether the fluorescent label degraded or somehow detached from the oligo, or whether the amplification was failing for some other reason.

Some paranoid biologists suggested that the companies producing the synthetic oligos have somehow altered the process in order to make the primers unstable, and that programmed obsolescence is part of a deviously clever money-making ploy. Although this is unlikely, the time frame for successful use of the oligos presents the companies with no reason to correct the problem. It is more probable, however, that the relaxed manner in which molecular biologists generally store their primer stocks has rendered them useless.

One possible cause of the failure is that the primers fall apart somehow or that trace contaminants left over from the chemical synthesis reactions change over time. This could cause degradation or modification of the synthetic DNA even at low temperatures, or inhibit the activity of Taq polymerase during the cycling program. For example, after the synthesis reactions, there could be a slow alteration of the oligo at the 3'-base, which may prohibit its extension by polymerase.

Although freezing and thawing has not been found to adversely affect the quality of double-stranded DNA samples as tested by visual inspection on ethidium-bromide-stained agarose gels [1], a plausible cause could be multiple freeze-thaw cycles. Roger Aeschbacher (aeschba@fmi.ch) suggested that some primer sets cause PCR failure by nonspecific priming because they are more likely to form primer dimers or intrastrand secondary structures, such as hairpins, only after undergoing multiple cycles of warming and slow freezing. This would not affect all sets of primers in the same way, explaining why some may require fewer freeze-thaw cycles to become nonfunctional in PCR relatively quickly, whilst others remain functional. That is, some primers fail after a certain time and others purchased from the same company on the same date work fine.

Waxing lyrical about hot start PCR
Performing hot start PCR can sometimes correct the problem. Hot start PCR is a method that produces cleaner PCR products by bringing the temperature of the template DNA and primer mix above the threshold of nonspecific annealing before the first enzymatic extension by the thermostable polymerase. This is accomplished by withholding a critical component from the reaction mixture, and is usually performed by separating the template and primers from the buffer components and thermostable polymerase with a physical barrier of wax. The wax melts between 60 degress C and 80 degress C, and the separated components mix by convection. Denaturation of template occurs and subsequent cycling begins [2,3].

The use of wax for hot start PCR has been a hot topic on the net. Instead of buying pre-fabricated wax nuggets for PCR, many netters have been making their own from blocks of low-melting-temperature paraffin, canning wax or Paraplast[R] medium, normally used for embedding pathological tissue specimens [4]. Others have invented their own recipes, mixing anything from 10% to 80% v/v mineral oil with the wax to soften it into more malleable lumps.

One method for creating beads the correct size to fit a PCR tube is to melt a chunk of wax at 80 degrees C and pipet 50-100 ul aliquots onto a sheet of aluminum foil or into the individual wells of a 96-well round-bottom microtiter plate (the latter gives uniformity to the shape of the pellets). After cooling, the foil/dish can be jarred upside down onto a piece of sterile filter paper on the benchtop to dislodge the nuggets, and a sterile syringe needle may be used to `prick up' the beads, which should be stored in a sterile container until used. One other useful method is to create a grease barrier, instead of a wax barrier, by dispensing generic petroleum jelly from a syringe fitted with an 18-gauge needle [5].

Protecting your primers
While hot start PCR may help reduce nonspecific annealing to template DNA for each individual problem primer set, this would be impractical on a larger scale. A more permanent fix could be to denature the bad primer stock at 95 degrees C to 100 degrees C for 5 min and then quickly cool it in a dry ice/ethanol bath or liquid nitrogen. This snap cooling would prevent any dimer formation and possibly restore the stock to its former condition.

Although it is still not clear that the primers went belly up because of the freezing and thawing cycles, one recommendation for protecting primer stocks may be to avoid temperature fluctuations by aliquoting out dilutions of the concentrated primers and storing them at 4 degrees C. Only thawing those to be used during the course of one or two experiments could save the original stock from an untimely demise. This would also be an intelligent prophylactic maneuver to avoid the risk of introducing nucleases from ungloved fingers, or other contaminants, into the undiluted primer stocks. A more long-term solution would be to store the primers at -70 degrees C as lyophilized aliquots.

References

[1] Anonymous (1983) BRL Focus 5(2), 10

[2] D'Aquila, R. T. et al. (1991) Nucleic Acids Res. 19, 3749

[3] Chou, Q. et al. (1992) Nucleic Acids Res. 20, 1717-1723

[4] Wainwright, L. A. and Seifert, H. S. (1993) BioTechniques 14, 34-36

[5] Horton, R. M., Hoppe, B. L. and Conti-Tronconi, B. M. (1994) BioTechniques 16, 42-43

Any statements made by the author are not meant to advocate the use of a particular commercial product or endorse any company. All opinions are those of the author and do not reflect the opinion of the National Cancer Institute or the National Institutes of Health.

Copyright: This manuscript is not copyrighted by Elsevier Publishing Company. However, you may not reproduce any portion for resale or edit the text for redistribution, sale, or otherwise without written permission from the author.

You found this at the World Wide Web (WWW) Uniform Resource Locator (URL):

ftp://ftp.ncifcrf.gov/pub/methods/TIBS/jan95.txt

Any reference to this column must be cited as the following published article:

@article{Hengen1995Jantibs,
author = "P. N. Hengen",
title = "Methods and reagents - Wayward {PCR} primers",
journal = "Trends in Biochemical Sciences",
volume = "20",
number = "1",
pages = "42-44",
month = "January",
year = "1995"}

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* Paul N. Hengen, Ph.D.                           /--------------------------/*
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