Hi Emma,

Your universal bacterial primers are not amplifying non-specifically, they are amplifying residual bacterial DNA present in your reagents, most likely coming from your Taq DNA pol. As there will normally only be approx 1-10 copies of the contaminating 16S gene in the volume of taq you use, you will only see the band coming up after 30 cycles.

The contamination in the Taq can either be from the bacterium (usually E. coli) used to make it or from waterborne bacteria from the water used in the Taq purification process. One way to find out what the contamination is is to sequence it.

Can I ask what is your "blank". How is it different to your negative control? If it has no PCR reagents in it how would you expect to see a PCR product in it?

No matter how careful you, are you will get amplification in your blank if you perform enough cycles using universal bacterial primers and your Taq is not DNA free. One company that sells "DNA free" Taq is Molzym but having spoken to some users of this enzyme, while it is better than most Taq on the market (in terms of contamination) you can still have problems.

My advice is to keep your cycles at 30 and continue with your experiments.

Also, you should start using real-time PCR instead of conventional!!

Hope this helps.

J.

Depend of the localization of the band, if you send a image of the electrophoresis should i help you

Maybe you had a contamination in your reagents.

Did it happen several times, using the same reagents?

If you loaded your product next to your negative control, it could have been as little as a tiny bit of spill over/leakage into the negative control lane, this would be my first guess as it's very easy to do, especially if your pipettes have pretty stiff plungers.

If your product is small (50-100bp), the annealing temp is low, and you have some strange secondary structures/binding sites predicted (IDT or wherever you get your primers from should have software to predict these things) then primer dimers could be possible, but it's a small chance it'd end up where your product is.

If you have a second negative control you could use, I'd sterilize all equipment/benches/tools related to your PCR setup and re-try with the new negative control and the old negative control. It could be as simple as a pipette having a tiny drop of template or product DNA in it prior to the PCR reaction. It's good to have sterile and dedicated PCR benches that are cleaned often and well for best results. Best of luck!

If the band is the same size, it is possible that your blank (presumably water) is contaminant free but that one of the components in your negative control is contaminated. Some Taq will do this so I recommend that you test each of the components in your negative control.

I think it could be a non-specific amplification as the blank is fine. I would also suggest to run your controls (both negative and blank-30 and 35 cycles) once again and load them by leaving one well between them to rule out the spill over.

Thanks for all of the suggestions. I initially thought it could be spill over from gel loading so I did re-run the negatives for the 30 and 35 cycle PCR, leaving gaps between them to avoid overspill and still the 35 cycle PCR negative has a faint band whilst the 30 cycle negative is completely clear.

I had the same problem with some of my PCRs. One of the problems I discovered is that sometimes your enzyme could have some DNA contamination in the solution. As far as I understand, most Taq is produced using E coli, and there is always some slight contamination with E coli DNA. Because it is a small amount, you won't see it until you start doing lots of cycles.

I would try getting an enzyme that has been purified. I use AmpliTaq Gold Low DNA for my higher cycle PCRs, and then the cheaper Taq for regular 25-30 cycle PCRs. It has worked pretty well. I also try to keep my area clean, wear a clean lab coat, and do all the other "clean PCR" stuff.

It's interesting that you have a clear negative control at 30 cycles and a faint band at 35 cycle. I still think you have little contaminated reagents and last 5 cycles are needed to let you see a band.

Hi everyone,

Thank you for all your comments/suggestions. I appreciate how difficult 16S bacterial PCRs can be as there are so many potential ways in which contamination can creep in!

I did use fresh water, and UV'd and cleaned all pipettes and equipment I used as well as using a fresh aliquot of Bioline MyTaq which is an all-in-one master mix that contains a high quality polymerase and is designed to produce high yields in fast reaction times. Hence, this is where my thinking came from about non-specific amplification as the PCR program for the primers I used is quite extensive and therefore usually works well on difficult to amplify samples, but the PCR's I mention here were just of bacterial DNA isolates being amplified for use as positive controls.

My plan is to make sure everything is clean again and run my next PCR again at 30 and 35 cycles and see if the same phenomenon occurs as then I think it is linked to non-specific amplification or contamination from the Taq due to a high number of PCR cycles and if I have cleaned everything and used fresh aliquots of everything, not removing the contamination source and/or contaminating the same 1 sample out of 46 seems unlikely!

Aliquot the nuclease-free water into small volumes after exposing the tubes to UV and expose the aliquots to UV while keeping the caps open. Also expose the PCR tubes similarly and run your PCR again. This has worked out in our case during qPCR. Check out the chances of contamination for each and every PCR component.

Thanks for the suggestions. I will take them on-board.

I was wondering what peoples' opinions would be on me taking forward the PCR products from the 30 cycle PCR where the PCR negative controls and the blanks were clear? I appreciate the comment that there could be contamination that you cannot see without a higher number of cycles, but it strikes me that with general bacterial PCRs, once you start going beyond 30-35 cycles, you start to get non-specific amplification? I previously did a nested PCR using PCR product from a first round PCR as the template for the second round and whenever I carried a negative control through both rounds, the negative control was clear in the first round and contaminated in the second round regardless of clean-up approach.

Hi Emma,

Your universal bacterial primers are not amplifying non-specifically, they are amplifying residual bacterial DNA present in your reagents, most likely coming from your Taq DNA pol. As there will normally only be approx 1-10 copies of the contaminating 16S gene in the volume of taq you use, you will only see the band coming up after 30 cycles.

The contamination in the Taq can either be from the bacterium (usually E. coli) used to make it or from waterborne bacteria from the water used in the Taq purification process. One way to find out what the contamination is is to sequence it.

Can I ask what is your "blank". How is it different to your negative control? If it has no PCR reagents in it how would you expect to see a PCR product in it?

No matter how careful you, are you will get amplification in your blank if you perform enough cycles using universal bacterial primers and your Taq is not DNA free. One company that sells "DNA free" Taq is Molzym but having spoken to some users of this enzyme, while it is better than most Taq on the market (in terms of contamination) you can still have problems.

My advice is to keep your cycles at 30 and continue with your experiments.

Also, you should start using real-time PCR instead of conventional!!

Hope this helps.

J.

Hi Justin,

Thanks for your answer. I will have a look into other Taq polymerases, but it is frustrating as I have changed Taq before and this always seems to come up as a problem! The Bioline MyTaq has worked well so far for me because I have difficult to amplify samples which it does a good job of amplifying in 30-35 cycles and now I feel anything more than 30 cycles is causing residual bacterial DNA amplification so 30 cycles has to be the limit, but I am confused as I have previously used these primers and Taq with 35 cycles and seen no contamination.

To clarify, the blank was just the nuclease-free UV'd water that has been used to create the master mix and used in the negative control as a substitute for any template. I included it to try and rule out any external contamination from the lab environment as opposed to from within the PCR.

Hi Emma,

Different batches of Taq will have different levels of contamination. One batch might produce a band after 35 cycles , another at 30 and another at 40.

Also, the more you run a particular PCR in the lab, the more likely you are to get residual PCR product contamination in your reagents.

Not easy I am afraid, but everybody has the same problems you have encountered.

J.

Hai Emma,

It is always advisable to get PCR product in minimal cycles unless there is a need for more number of cycles. In your case you have ruled most of the possible reasons for cross contamination but whether you changed the stock primer dilution or the working primer dilution because contamination may be occurred in that level also. For me it happened once and I've changed the stock primer solutions and my negative became free of contaminated product.

Hi Jayakar,

Thanks for your comment. Yes, I will be using fresh stock and making fresh working primer solution from this in my next PCR!

Contamination in 16S bacterial PCRs is a common problem for many, so it's great to be having this discussion and sharing ideas with everyone!

Most commercially available Taq would come from a cloning product in E. coli. You can try using native Taq, Taq that were produced from the original Thermus aquaticus, but still no guarantee that u will have no band after 35 cycles since its also depends on how the sequence of your primers match the DNA sequence from Thermus aquaticus itself.

Primary source of contamination would be the person doing the experiments since our body are actually teeming with comensale bacteria they tend to shed all over the place, so its just a matter of time before that one or 2 bacteria are lucky enough to contaminate your fresh PCR reagents or clean working space (unless u can isolate everything like in a glove box)

Hey Emma,

most of things have been covered in the discussion. Just one more suggestion. To rule out PCR carry-over contamination you could also try using Uracil DNA glycosylase (UDG) in your PCRs. That way you can rule out amplicon contamination. All the best. I used to consider my day as a good one if i didn't get a band in my NC when i was doing 16s PCRs. Hope you have a good day!

Dear Emma,

35 cycles is quite excessive, and therefore any minimal background contamination will be amplified in your samples. As others have recommended, I suggest aliquoting out your reagents. I had problems with 16S contamination in a previous lab (even at 25 cycles), and found that the only way to combat this was UV my water.

Good luck.

the result will be also primer-dependent, not all universal 16S rDNA will produce the same "contamination". Some not et all. probably related to primer efficacy and self-priming potential. You can expose reagents to heat-labile DNAse I, or UV and "decontaminate" before use. We noticed huge difference in primers ability to detect "contamination". what is your objective at Ct>35?

Thanks for your comment Ivan. I can appreciate that different primers may vary in their ability to detect contamination. My objective in using 35 cycles was to improve amplification as the samples in question were of a low yield. When I repeated these PCRs with a different batch of master mix I did not detect any contamination even at 35 cycles, but in general now I stick to no more than 30 cycles unless amplification is poor at 30.