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PCR Troubleshooting Checklist


Routine PCR?


Let’s be honest, there’s no such thing. Even with the simplest PCR reaction things can go wrong, so you need to have a good checklist of ideas for PCR troubleshooting and rectifying the problem. Today I have brainstormed all of the ways I can think of to approach problems with standard PCR reactions.

I’ve inevitably missed some things out, so please chip in if you can think of anything else to add. I will add your ideas to the list to make it a resource we can all refer to.

No Amplification .

1. Try the reaction again, you may have left something out.

2. Check that the polymerase buffer has been fully thawed and mixed thoroughly.

3. Check that the primers have been diluted to the correct concentration.

4. Make up a new dNTP solution. dNTPs can be destroyed by repeated freeze-thaw cycles.

5. Re-make the template DNA, especially if you are working with genomic DNA. Old stocks may be degraded or sheared.

6. Use a different polymerase. If an amplification is problematic with a proofreading polymerase, I often try using good old Taq and this often solves the problem. Remember to sequence the insert though – if you are lucky there won’t be any significant mutations and you can use the insert as it is. Otherwise, re-do the PCR with a mixture of Taq and 1/10th concentration of your proof-reading enzyme and you should still get the same amplification with a lower error rate.

7. Change the annealing temperature. If the annealing temperature is too high, you obviously won’t get any priming at your desired sequence. On the other hand, an annealing temperature that is too low can result in such non-specific priming that don’t allow specific bands to arise. The best way to find the optimum temperature is to use a gradient cycler and test a range from the lowest primer Tm to 10ºC below in 1ºC increments.

8. Try reactions with varying template concentrations. Your template concentration may be too low or the concentration of impurities in your prep may be too high. Try 5-10 parallel reactions with concentrations from 10 to 200 ng in a 50 microlitre reaction.

9. Check the cycler – are the temperatures and times as you expect?

10. Try the reaction in another cycler – the calibration of the one you are using may be off.

11. Try an additive. I find that DMSO is especially useful in problematic amplifications.

12. Redesign the primers – try to stick to the guidelines as closely as possible.


Non-specific Band Amplification .

13. Re-do the reaction with a negative control (no template). The non-specific bands could be from contamination of one of your stocks with foreign DNA (probably yours!). If this is a problem, use new stocks, always use autoclaved PCR vials and wear gloves and a lab coat.

14. Increase the annealing temperature. Better yet, use a gradient PCR machine (see 7.)

15. Redesign the primers and make the 3′ longer. The extra bands may be from similar sequences to your target. Increasing the primer length will make them more specific for your target.

16. Increase annealing time if the non-specific products are shorter than your target. If they are longer than you target, reduce the annealing time.

17. Use less DNA template.

18. Try touch-down PCR.


Weak Amplification of your Target .

19. Reduce the annealing temperature.

20. Increase the annealing time.

21. Increase primer, template and/or polymerase concentrations.

22. Try touch-down PCR.

23. Increase the number of cycles.

24. Try an additive.

25. Clean up the isolated target and use it as the template in a new reaction.

…now, what have I missed out?


Originally published on January 23, 2008.  Revised and updated on July 13, 2016.



1) elahe on October 31, 2018 at 1:58 pm .

in have 1 big problem in may work
my positive control not band however i checked my Components master mix
this dna of control Worked before
i checked master mix mgcl dntp and
Amount of dna
i realy confuse

2) Amanda on July 24, 2018 at 8:21 pm .

Also, watch out for MgCl concentrations: higher concentrations of Mg+ ions can increase polymerase activity, but at the expense of specificity, so could result in more nonspecific amplification. The usual way to overcome that if you can’t reduce MgCl (a lot of high fidelity/high activity polymerase master mixes have high MgCl) is to increase annealing temps.


Roxanne on August 7, 2016 at 7:12 pm .
There are 3 things that I would add to this already thorough list:
1) If your amplicon is really weak or not showing up at all, supplementing with MgCl2 helps a lot. Look at your Taq’s product sheet to figure out how much they suggest you add for troubleshooting.
2) Use filter tips! It might sound wasteful, but I’ve seen the difference they make with my own eyes.
3) Water!! That’s the first thing I change when a PCR doesn’t work. Once I used freshly autoclaved ddH2O side by side with “store-bought” nuclease-free water in a particularly tricky PCR and only the ladder reaction gave me my amplicon. You never know what may be in your water.

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Suzanne Kennedy on August 7, 2010 at 8:46 pm .
Are you using different pipettors for the PCR set up than the ones you use for everything else?
If you have a hood with a UV light (in another room) you can set up PCR in there.

We first clean our hood with 10% bleach including the outside of tip boxes. Then we wipe everything down with 70% ethanol.
Then we put everything in there we will use. We also use disposable arm protectors so we put those in the hood. We put the UV light on and irradiate the hood and the outside of pipettes, bottles, tips, and the PCR tubes and tube racks.

Now set up your PCR. Put the matermix into your tubes and set up the water controls. We add the genomic DNA template last and outside of this hood after the water controls are all closed and done.

This should do it.
When we are finished, we wipe down the PCR hood again with bleach and ethanol and UV it to decontaminate.

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Tamara Miller on August 7, 2010 at 12:24 am .
I am having problems with amplification in negative and water controls. The product always ends up the exact size of my expected product. I’ve tried almost everything I can think of to solve this problem, including buying all new reagents, using filtered tips and pre-packaged tubes, re-ordering primers, used a thermocycler in another lab etc. etc. Next I will try another run using only water controls and set up the reaction under a hood.

Truly people, I am stumped, desperate, and in great need of help.

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Mostafa Najafi on September 13, 2016 at 11:32 am .
Hi lm mostafa

I have this problem too.

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Israa on December 19, 2016 at 2:21 am .
Hi Tamara
I have the same problem like you and I do not know how to sort out I,m desperately strugle

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Nafiisah Chotun on July 5, 2010 at 7:28 am .
What about reducing the extension time if your PCR product is a short one (300bp)? Would that work?

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Poonam Pawar on July 1, 2010 at 6:34 am .
I had encountered this really weird problem of getting amplification in the negative control and now i suppose ur idea of having the concentration of impurities in prep may be too high is right n i will surely Try 5-10 parallel reactions with concentrations from 10 to 200 ng in a 50 microlitre reaction as u have suggested.

this is been real help n i will surely try abnd let u know if it helps.

thanks nick.

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vasuprada on April 28, 2009 at 4:00 am .
Any thoughts on MgCl2 concentration when no amplicons are observed?..coz i get postive control bands and one target gene amplicon from a total of 8 genes, so i know my reagents are good. Any help is surely welcome! thanks in advance!

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RKSmith on September 23, 2008 at 7:40 am .
I sometimes use a double positive control which is a positive control with pure DNA then a second positive with pure DNA PLUS aliqout of components used to isolate the real samples (i.e. same stuff used for sample purification) to detect an inhibitor situation. If the pure DNA works and the seoond positive with components used for isolating your samples doesn’t then you know you have an inhibitor prolem (try Bovine serum albumin to correct this or another clean up to purify template better).

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Jacqueline Lopez on February 8, 2008 at 8:54 pm .
I have found that dNTPs that are diluted in water are more sensitive to freeze-thaw cycles than those in 1XTE buffer.

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maximilianh on January 23, 2008 at 6:54 pm .
I’ve heard that NTPs are sensitive to freeze-thaw cycles… However, my whole lab is happily freezing-thawing them for months now and it doesn’t seem to be a big problem…

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Ben T. on July 27, 2016 at 10:22 am .
This “freeze-thaw” thing drives me crazy. Enzymes are sensitive to freezing and thawing; small molecules like dNTPs are not. They DO break down — to nucleotide diphosphates and monophosphates, mostly — over time, and the rate of breakdown is proportional to temperature, as is every chemical reaction. So the longer your aliquots have sat on ice, instead of frozen at -20, the more degraded they will get. But how many times you freeze and thaw them is totally irrelevant.

In my experience, individual dNTP aliquots can be used dozens of times without getting degraded enough to interfere with PCR efficiency. dNTPs are fairly chemically stable.

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Dr Amanda Welch on July 27, 2016 at 5:50 pm .
I would agree with you on that they’re very stable. However, they do breakdown eventually and freezing and thawing does increase the rate that they breakdown. So, it’s worth making up a new solution as part of your troubleshooting. Although, I might be a bit biased, because I’m speaking as someone who had a PCR not work because of degraded dNTPs. 😀

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Chad on January 23, 2008 at 3:08 pm .
Geez, I can’t think of anything to add off the top of my head. That was a pretty thorough list.

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