We had the same problem, but finally we found out that some types of polymerase cointain rests of E.coli. Sometimes happens that certain LOTs of polymerase have been contaminated in the production process, but other LOT work OK. Try another type of polymerase! If you want to be sure that your laboratory (gloves, tubes, pipetes) is not contaminated, clone and sequence your PCR contamination product. If you get some Pseudomonas sequences, the problem is the sterility of your lab. If you get some E.coli, the cause of the contamination is the polymerase itself.

Hi Liz! I will refer you to this thread: https://www.researchgate.net/post/We_are_frequently_seeing_reactions_in_our_PCR_negative_control_water_added_no_DNA_when_we_do_RLBs_any_help?ev=home_person_like_comment_question_target

The only source of contamination is your Taq polymerase which is purified either from E.coli or other bacterial sources. There is always bacterial DNA remnant from the commercial preparation. If you produce yourself Taq you will realize it. So, my advice is try to incubate the Taq polymerase with DNase I in Taq buffer supplemented with MgCl2 (1.5 mM will be fine) and 0.5 mM CaCl2, heat-inactivate it and then add your primers, dNTPS and the samples, and proceed.

We had the same problem, but finally we found out that some types of polymerase cointain rests of E.coli. Sometimes happens that certain LOTs of polymerase have been contaminated in the production process, but other LOT work OK. Try another type of polymerase! If you want to be sure that your laboratory (gloves, tubes, pipetes) is not contaminated, clone and sequence your PCR contamination product. If you get some Pseudomonas sequences, the problem is the sterility of your lab. If you get some E.coli, the cause of the contamination is the polymerase itself.

what do you mean about negative control is that mean non template control or using negative sample, my advice to you is

1- use non template control to check the contamination (if you don't use it )

2- if you use negative sample check primer mismatching and another negative sample

3- your DNA sample could be contaminated so try to re isolate all off your samples

4- finally if you decrease the number of cycles you will get rid of the faint band but it will still be there if you increase the number of cycles

Hope it help you

I understood she UV all the sample mix except the Taq and the DNA. In principle, the negative control should not give her any band in those conditions. Because that means she got the contamination at time of adding the Taq pol. Again,in my opinion 99.9999% chance the suspect is Taq I think from what she has described, she is working under super clean environment and with high precaution than most of the labs do.

I agree with Maria. She should sequence those bands to trace back the source of contamination.

The bands in your negative controls are of the same or different size than the expected product?

This is important to understand if you have unspecific or specific amplification. In case the negative control is a non-DNA control as Yasser pointed out you shouldn´t have any band and then it is a contamination of reagents. If your negative control is a negative DNA and you have an unspecific product (different size) maybe your primers have little identity with another sequence in your sample.

Another important point, even if it doesn´t seem to be the case here, is to maintain pre- and post-PCR processing of samples physically separated as it is very common that contaminations (specific product in this case) come from the already amplified samples. If, for example, you use your dedicated pipetes to load the gel to check the products there you have your contamination source.

I came in to say check your polymerase as well. Very often, it isn't completely purified and has carryover DNA from the bacteria used to make it.

also check primer dimer properties

quantity and quality of DNA

optimize PCR conditions

Maybe you should use the Taq polmeraze and buffer from the same producer, or if you already did that, try with the newer ammonium buffer, it sometimes fails when stays long on 4 Centigrades. My team and me were in the same problem and we were suggested also to apply Bovine Serum Albumin (BSA) in the amplification mixture and that seemed to work out, we are still checking that.

I think sometimes it happens people do not understand the issue because of the way it is formulated. That’s what I have understood. Liz is getting amplification in its negative control sample. That is (water + PCR buffer+ dNTPs+ oligo+ Taq). She Subjects all the sample mix to 5 min UV treatment before she add the Taq polymerase. In principle, even if she had any contaminating DNA it should give absolutely no signal at all.

no its not like that she has mentioned that she exposed everthing to UV except reagents and DNA

In this case, I think she has to do that to test her negative control and then report back! That is to UV for 5 min the PCR mix (water + PCR buffer+ dNTPs+ oligos) and then add Taq. She can split just before UV treatment: some samples exposed to UV and others not exposed to compare.

UV gives only superficial sterilization... it won´t work on the liquid contents of a microcentrifuge tube that is plastic (UV doesn´t transverse this material).

Hi María: Very longtime ago, I was taught to get rid of contamination using UV and did rise the same argument as you but that maybe only for old thick old PCR tubes. UV irradiation spectrum is relatively broad and not all plastics are made from the same polymers. At 254 nm it does traverse very efficiently the thin transparent 0.05 and 0.02 ml PCR tubes. If one is using microplates and has access to a UV-crosslinker chamber that makes it even easier. I have used both the transilluminator and UV-crosslinking device (Startalinker) with good results. I must say I was including even dNTPs, which may reduce the efficiency of UV on the unwanted contaminating DNA. I even think that using the UV in the UV-hood should work.

So, anyway I think Liz must have a look to this work on how to get rid of contamination. . (http://www.plosone.org/article/info%3Adoi%2F10.1371%2Fjournal.pone.0013042). There are more refs on the application of UV to PCR to reduce contamination.

Aswering to Abdelhalim I must say that I do not have experience in decontaminating solutions with UV-crosslinker but standard UV lamps as those in laminar flow hoods do not have high penetration into liquids and are far away from the work surface. Why do you say you had good results? did you introduce a contamination on purpose and then measure that it was eliminated?

Indeed the article by Champlot et al. published in PLoS ONE that you suggested to Liz is really good and explains how UV irradiation must be used in a way that it effectively decontaminates reagents for PCR. It also clearly shows that it is not straight forward that just irradiating with UV light would make it. You can decontaminate certain reagents but not others, at a very close distance from the bulb, making sure that the plastic on your tubes is not absorbing UV light, measuring the exposure time, etc., etc. and combine it with other treatments.

Maria using UV with specific WL is some time creates problem like mutation, i.e. base changes in DNA so using UV thats why always risky and I am not sure whether you can use that technology for decontamination. Elizabeth you can use Millipore columns for DNA or PCr product purification which are robust and popular or can read Forensic DNA articles on that. I was in that field Forensic DNA, and as you know in Forensic, the major issue is contamination and tissue degradation.

regards

Dr.Prabal

Exactly Prabal, I think the same way.

Maria and Prabal: I think what one wants is to neutralize the decontaminating DNA so as it is not used as substrate by Taq polymerase. UV-induced mutations need a DNA repair pathway that operates into a live cell. This has nothing to do with the issue.

Maria: No I didn’t introduce any defined fragment. Yet, I always solved my contamination problems using UV since I was taught. Sometimes sequencing the products in the negative control gives you an idea on the source. I did it just once or twice. The UV transilluminator also gave me good results. So, I always UV my PCR mix just before adding the polymerase. As you know in most labs PCR is performed on the bench at least for most applications. There are plasmids, specific PCR fragments, cDNAs, etc… everywhere in the lab.

I know of people who have UV-irradiated cells in their culture plate (in PBS buffer with less volume) using the UV-lamp in the laminar flow hood. They were a bit inventive sometimes by lifting the plates to a reasonable distance from the lamp. I know it is not the proper way to do it (irreproducible) but it activates DNA repair pathway as demonstrated by specific markers that all what I can tell you. Anyway, that is not the question here.

Dear Elizabeth,

here's a list of suggestions:

- wipe the whole hood with ethanol70% and turn on UV lights for at least 15 minutes (put the RNase free water eppendorf in the hood as well);

- very important: re-order primers and pay extreme attention to contamination while eluting and diluting;

- re-order the taq, buffer, mg, BSA...;

- reduce mg content to improve specificity (be sure it is enough to amplify your fragment, though).

ciao

Andrea

The best way to get rid of contamination is using 10% bleach. See: http://bitesizebio.com/articles/pcr-the-right-way-to-decontaminate-and-eliminate-false-positives/

and the paper it references: http://www.ncbi.nlm.nih.gov/pubmed/1571142

Chuck out your old reagents and use fresh stock of everything to ensure you've not contaminated them. If you are trying to amplify E coli then you could be getting contamination from traces of E coli gDNA in the Taq you use (assuming that this is what was used to make the Taq). Talk to your supplier and see if they have any Taq they can guarantee as being E coli DNA-free.

I concur – I suspect contaminated reagents – fresh stocks of everything including primers and all from alternative suppliers. A fresh lot from the same supplier can still give the same problem. Alternative primer design on top of this would be even better – since you’re ordering new primers anyway.

We aliquot everything into small tubes, and store it all at -20, and thaw small amounts when we need them.

I will try a new tube of Taq polymerase, and new reagents.

The amplification I am getting in the negative control is a faint band of the same size as my product. I wish I could reduce the number of cycles, but the point is to amplify those small amounts of DNA.

I have tried BSA in the past, but did not see much improvement.

Thank you so much for all your advice. I will read those articles, and give it another try.

There's a more recent, very comprehensive, article on contamination and clean-up here: http://www.plosone.org/article/info:doi/10.1371/journal.pone.0013042?imageURI=info:doi/10.1371/journal.pone.0013042.t001

Faced a similar probs but my issue i vortex-ed my epp. After adding. Then only i realized i needed to invert the epp.

Sometimes the contamination problems are located inside the micropipets (micropipettes), you should try, simple by interchanging pipets with some mate which does not work with E coli.

On the other hand, this type of problem is frequent by using some types of Taq Polymerase, assay several trademarks, Roche taq woks well;

have you considered that you do not have contamination but instead have some non-specific amplification going on in the negative control? I assume the negative control has some source of DNA. It could be that your primers are binding to DNA sequence in the negative control that some homology but is not your intended target. It could also be that your primers make primer dimer product even in the absence of any target. You should not need 40 cycles to pick up even small amounts of your target. perhaps cut back your cycles to 35-37 cycles and/or try a little but higher elongation/annealing temperatures to see if you get rid of non-specific amplification.

Carry out PCR reaction using another primers to be confirmed from the purity of reagents, gloves, pipetes...etc. Also, try to modify PCR conditions: denaturation, annealing, extension (temp. and duration) and cycles number. Good luck

You can try a nested PCR instead of 40 cycles. If you still obtain a band then you can be sure you have contamination and it is from the same origin of your sample.

There are several people above with good answers. First, ethanol doesn't get rid of DNA... use bleach and/or UV irradiation and continue working in a HEPA filtered hood. Second, the bacterial polymerases we all use end up with residual bacterial DNA after the polymerase is purified away from the bulk culture during production. This may vary from lot to lot and company to company, but if you're trying to amplify low levels of target DNA, chance are that you'd see it. Several companies offer low bacterial DNA qualified polymerases (I don't know who off the top of my head), but beware that it will typically add extra expense.

What is your negative control? If you have some DNA as your negative control change the DNA. Otherwise buy polymerase from a different vendor. If those do not work, then take a vacation and come back start afresh. Your boss will understand!

UV decontamination is not an efficient way. Read this article on DNA decontamination:

http://www.plosone.org/article/info%3Adoi%2F10.1371%2Fjournal.pone.0013042#s2

have had the same as Maria. Change all your reagents and try a different Taq. I would also proceed with Sequencing to see whether the products are the same. You may also need to re-design primers outside the current set just for PCR. and use the internals to sequence with

All answers above are great. Try to include a no template negative control (i.e. master mix only plus your PCR water). If you did this and still got a positive negative control, then use a new set of all components including water and DNA. However, note that your PCR product is one of the most stable contaminants for your next PCR.

I too faced a similar problem only to discover that the primer set got contaminated. My PCR worked fine after changing the primers.

Hello ...I agree with Mária Džunková answer,Please check your DNA concentartion for PCR amplification.What is the Nanodrop ratio your using for PCR ammplificaton.

Hi. The optimization of the PCR conditions (cycles, temperature and time) seems to be a good idea. Maybe the addition of components like BSA etc will reduce the background noise in your negative control. You can also try to change the concentrations of Potassium and Ammonium ion to achieve a more stringent amplification.

Good luck.

All answers above probably solved your problem. I would suggest to use filter tips and different polymerases.

I would like to suggest you that plz check your primer and change your Taq at once. Most probably this is due to taq and reduce your PCR cycle. May be your taq is not effective till 40 cycle. After that keep a temperature gradient to find out the best annealing temperature

I suggest the following to overcome your difficulties:

1. Wash micropipette barrels in 3% sodium hypochlorite solution (but contact the manufacturer of the micropipette to ensure that this is suitable for your brand of micropipette).

2. Rinse the pipette barrels in Ultra-pure water from Sigma Chemical company.

3. Dry the pipette barrels upright in a laminar air flow works station.

4, Buy a new box of sterile filter tips suitable for PCR applications and place in the laminar air flow work station.

5. If you are using PCR tubes to carry out the reactions please use an unopened bag of tubes and handle with sterile gloves inside a laminar air flow work station.

6. Purchase new reagents, dNTPs, oligonucleotide primers, sterile ultra-pure water (sigma chemical company), 10 X PCR buffers, 25 mM MgCl2 stock solutions (for PCR applications) and Taq polymerase and taq dilution buffer and PCR oil (if using PCR oil to prevent evaporation within tubes)

7. Place all the materials needed except the template material into one dedicated laminar air flow work station. Manipulate all reagents, enzymes, (PCR oil if using PCR oil) using filter micropipette tips in the laminar air flow work station. Retain three negative control PCR tubes that will have no template material added in this laminar air flow work station.

9. Move the remaining "test" PCR tubes (now containing all ingredients except template DNA) to a second dedicated laminar air flow work station preferably in a different room. Add the template material to these tubes using a dedicated micropipette.

10. Place all tubes 3 negative controls and remaining PCR "test" tubes into the PCR machine and cycle for 30 to forty amplification cycles.

11. Prepare the agarose gel containing Sybr Green stain for visualisation and analysis of the PCR reaction products.

12. Load the DNA from the negative controls into the agarose wells first of all using a clean dry micropipette with sterile filter tips.

13. Leave an empty sample well in the agarose gel to avoid spillover of samples into adjacent wells.

14. Using a different micropipette that has been cleaned and dried and fitted with micropipette filter tips load the PCR "test" samples into the agarose gel, again with gaps between sample wells, to ensure again no spill over contamination occurs when loading wells.

Dear, Wipe your hood and other things with DNA ZAP solution. this will surely help you.

What types of primer are you using, if its 16srRNA, it will amplify with most of microbial strain and the result you'll get a PCR product if there is tiny tiny contamination. Usually work with sterilized filtered tips. And use 10% bleach or any strong detergent instead of EtoH in which ethanol will precipitate any DNA residues 

Ethanol does not get rid of DNA, definitely use DNA zap. Also buy new pipettes and use them only for PCR. You should not use the same pipette which you have used for DNA extraction. Some DNA may have already got into the barrel of your pipette during extraction and it is impossible to clean it out.