Here is what you need to do if you work in a large group or lab dealing with too many biological materials

1) Use a clean seperate set of pipettes and use barrier tips for all work including stock reagent preparation

2) make ur own stock all all items including DNTPs, buffer, water and primer from the scratch(make small aliquots to prevent contamination and too many freeze thaw cycles.)

3) use a good source of water (distilled or milliq water is NO for PCR grade water. IF you must there are some precautions)

4) all plastic ware must be clean( we use fresh pcr grade tubes and open only in a laminar hood, and make batches in small containers.)

5) Maintain good practice of molecular biology lab, like correct ways to use gloves avoid aerosol contamination etc.

If you try to troubleshoot at the point where you are right now there are too many variables that will take up your time with no solid result, Hence save time and effort ..start fresh.

Best wishes

Hi Daniel,

Maybe the tubes are contaminated ? Try to use sterilized tips with filters, it may help!

Please check this.

http://www.ncbi.nlm.nih.gov/pubmed/16905333

Hi Daniel

I agree with Dawid, if you used new mix and water your contamination may be in the tips or pipettes. Once you clean them make sure that nobody use them for other experiments. Are you using your own DEPC water? if so you may need to prepare a new stock and prepare aliquots.

good luck

Hi Daniel,

What is the sizes of the pcr product in the negative control? Is it similar to your samples or smaller than 16S bands? As previously noted, contamination can come from unstereilized tips/tubes.

Hi there,

UV sterilise all plastic-wear, PCR buffer water for 20 minutes before making up your master mix. Always make your mater mix up in a PCR hood (turn the UV lamp on for 20 minutes before you start). As mentioned, always use new tips etc between master mixes.

Cheers

Yes I m also agreed with others but primer's purification grade and water purification grade can't be ignored and remaining is true as earlier researcher discussed.

All the best

I agree with Jack on the UV hood - where possible I also put my water in the UV hood too, regardless of whether it is brand new.

Is it possible to reduce the number of cycles in your PCR too? You can optimise with the least number of cycles necessary. I also found that changing to a pre-mixed taq mix helped too, rather than one I made up myself (I'm not sure if that's what you meant by "new Tag Master mixture") - fewer steps for contamination.

Good luck.

Hi,

contamination can be due to improper sterilization.but if the molecular weight of the contaminant is similar to the pcr product, then it is your handling problem. Autoclave all the reagents and apparatuses work inside the hood always.

Good luck

Hi,

if you made primer dilution with nuclease free water or not? if not then go for it, dissolved you primer pellets and dilution in nuclease free water in hood, always used autoclaved tips for PCR reaction; Good luck

use different sample amount, if it works good, if not then use different quantity of dntp as well as mgcl2 co-facters.

Do all your pipetting in a cell culture hood.

And always be super cautious with your autoclaved tips... even if there is a slight chance that it touched something from the bench, etc. throw it away and get a new one. Surprisingly I've seen people continue aliquoting after touching the bench, and then complaining of having contamination in their negative controls. You can never be too careful with contamination!

Make use of fresh sterilized stuff and in the aseptic zone and use gloves always.

Here is what you need to do if you work in a large group or lab dealing with too many biological materials

1) Use a clean seperate set of pipettes and use barrier tips for all work including stock reagent preparation

2) make ur own stock all all items including DNTPs, buffer, water and primer from the scratch(make small aliquots to prevent contamination and too many freeze thaw cycles.)

3) use a good source of water (distilled or milliq water is NO for PCR grade water. IF you must there are some precautions)

4) all plastic ware must be clean( we use fresh pcr grade tubes and open only in a laminar hood, and make batches in small containers.)

5) Maintain good practice of molecular biology lab, like correct ways to use gloves avoid aerosol contamination etc.

If you try to troubleshoot at the point where you are right now there are too many variables that will take up your time with no solid result, Hence save time and effort ..start fresh.

Best wishes

One issue that has not bee addressed it your polymerase. The are from purified from bacteria, and most will have contaminating bacterial RNA. We use 16S to confirm that our environmental extracts are amplifiable, and our water control is always expected to amplify, though to a lesser extent.

You can try to obtain a more purified Taq (or other polymerase), but that may not solve your problem.

Good luck.

We do 16s as a clinical service, and no matter what we do, the negative controls are always positive after about 32 cycles. There are traces of bacterial DNA everywhere! I tried UV decontamination of everything but it made no difference whatsoever. We make sure that we use all new disposable plasticware (tips and tubes etc), molecular grade water and reagents that we keep exclusively for 16s, where possible make everything single use aliquots, use a highly purified low DNA polymerase.

Hannah, I have the same experience. Running 32 cycles makes the negative control almost always a tiny band. Now I am using 30 cycles and the result looks good. I can hardly see any bands for negative control.

We do real time PCR so we can see the amplification curves and dissociation curves not just bands. We do 35 cycles but simply compare our samples with the negatives. Anything that amplifies two cycles earlier than the negatives is considered a true positive.

You can treat your PCR mixture - minus the sample DNA and primers, but with Taq polymerse and everything else - with DNase/RNase beforehand. Make sure you denature the DNase/RNase before adding any sample DNA or PCR primers. The Taq should not be negatively affected by this treatment.

James, DNase is easy to denature - simply heat the mixture to 95C. But how would you propose getting rid of the RNase? I suppose a sufficient amount of RNase inhibitor (RNasin, Promega) could be added, but I wouldn't have complete confidence in it.

Make single use aliquots of water for making your mastermix and store them and all other PCR reagents away from communal PCR stations or storage locations. It may help to also have a dedicated pipette set for master mix preparation as well as template addition. Change gloves liberally.

Then there is the issue of residual DNA contamination in the recombinant Taq Polymerases too. You may want to look into companies that make DNA-free recombinant Taq. Sigma sells MTP Taq DNA Polymerase (D7067), but even their product is "low contaminant", not DNA-free.

If your contaminating (NTC/negative control) Cq value is more than 6 cycles away from the Cq values in your samples, you have little to worry about during mathematical quantification. The contribution to signal from background room bacterial gDNA (always floating around) is negligible.

At what Cq does your contaminating signal come up? 30 or 32?

And your 16S signal comes up around 20 or less I would assume. If so, no worries, this is a hard problem to get rid of sometimes with 16S in bacterial qPCR and 18S with mammalian and plant qPCR when the rooms we must work in are not ideal (mostly due to air-flow/air vents/opening and closing doors, dust...)

Just something to keep in mind so you don't spend all of your time being too paranoid about every reagent and every pipet and every unclean co-worker ;]

gDNA contribution factor = Eamp^(Cq of sample - Cq of NTC rxn), and if it is assumed all reactions have Eamp of 2 (e.g., are 100% efficient), the gDNA contribution to final calculated relative quantity is only 1.5625%.

Contaminating RNA can be a problem if there is enough of it to sequester/bind one or both primers enough to take them away from the qPCR... since RNA:DNA hybrids bind more strongly than DNA:DNA hybrids, this can be a concern as James Borrone has alluded to.

@ Richard Pilon

Guess I should have been specific.

If you really are concerned about RNA (which should not necessarily be a problem) you can use:

RNase If from NEB, New England Biolabs: product M0423S or M0423L.

According to NEB, it can be denatured by incubation at temps at or greater than 70 C for 20 mins.

RNA should not be an issue though.

We have had similar and frustrating contamination issues with 8F x 515R (V1/2/3) combinations, despite working in different rooms, filtertips and dedicated pipettes, and brand-new batches of mastermix or water. When we switched to 338F x 906R (V3/4/5) (or 338F x 806R) PCRs the contamination problem was suddenly resolved. I guess there is just something that the 515R primer picked up.

As previously Paul A Bartley advised me: you can try use 70%-30% mixture of dNTP's/dUTP's. After that reagents can be decontaminated with the Ung enzyme.

If we have a problem with PCR contamination we are using blue, plastic autoclavable sticks from Robbins Scientific to open the tubes. And we treat everything what is possible with UV.

PCR diagnostic labs that 40+ cycles to detect rare DNA and RNAs like virus and parasites do one of two thing to remedy this problem. Use U instead of T in the PCR mastermix, and treat all PCR reactions with UNG before every PCR, This only works if you are setting up a new primer pair. Otherwise if you have already been using the primer pairs in the lab, make up your PCR mastermix including everything but template and the TAQ. You then spin the mix through a 30k molecular weight cutoff filter and freeze aliquots. This cutoff is between the primers and the pcr product. Even low level contaminated primers can be cleaned up this way. Before your PCR run, just add TAQ (be really careful with the TAQ, we added this to aliquots in another part of the building) and your template. This simple proceedure saved the company I was working for.

Sterilization is critical to diminish the contamination. Both UV and autoclave work well for general PCR. Preparing PCR matrix in another room might also help. However, when working with 16S universal primer, something else need to pay attention to. As mentioned above, Tag polymerase is a source of contamination because it contains trace of DNA from host bacteria that express DNA polymerase. The trace of DNA will cause contamination problem after running 32 cylces when the real PCR products usually get saturated but the trace DNA will be amplified dramatically. The simplest way to get rid of contamination is to run just 30 to 32 cycles while we still get enough PCR products for downstream processes.

We had a problem with one of our primers for another gene that we PCR before cloning, and couldn't understand what was the issue because we do PCR in the UV-hood and everything is treated. Then I noticed that our expected band was approx 100bp, and the primers are approx 25bp each. So I gave it a go, extracted the 'negative' band from the gel and sequenced it: it turned out it was just two primer-dimers attached together (therefore the same size of our band).

You may try to sequence your negative band and see if it's a real product. If it's just primers, then you can denature them for a 10min at 95C before your PCR and they should not give you problems.

I have a problem. Negative con trol was contanimate and it has size equal pcr product form bioinformatic data. I try to 10 times but it not ok all. May you explian abot method for solve a problem

Hello, Yijie,

You may find this article helpful: https://www.promega.com/-/media/files/resources/profiles-in-dna/802/identifying-and-preventing-dna-contamination-in-a-dna-typing-laboratory.pdf?la=en

Hope this helps.

Best,

Arizaldo

The main sources of PCR contamination in a laboratory are:

The last source of contamination, often called "carryover" contamination, is the most troublesome source.

One of the main approaches to control PCR contamination is to physically separate the working areas for:

· Preparation of the DNA template

· Setting up the PCR reaction mix

· Post PCR analysis

Many labs have a designated room for setting up the PCR reaction mix (pre-PCR room) and a separate lab for analysing the PCR products (post-PCR room). No items should enter the pre-PCR room from the post-PCR room and a separate clean lab coat should be kept for use in the pre-PCR room. if you don’t have a pre-PCR room use a PCR hood or cell culture hood or a different part of the lab to set up the PCRs.

• Use sterile techniques & always wear fresh disposable close fitting gloves when working in the pre-PCR clean room / area.

• Before you start clean bench surfaces and pipettes with a DNA decontamination reagent eg DNA Away

• Aliquot all reagents used for PCR into smaller volumes for storage.

• Use a dedicated set of pipettes for aliquoting PCR reagents and setting up the PCR mix.

• Use aerosol-resistant filter pipette tips or positive-displacement tips.

• Always Use PCR grade water throughout.

• Never put hands inside PCR tube bags. Always pour out the PCR tubes on a clean tissue placed on a tray or surface that has been wiped with a DNA contamination reagent.

• Keep fingers away from the top of the PCR tubes as much as possible. Open and close all tubes carefully making sure you don’t touch the inside of the caps or rim of the tubes.

• Handle PCR tubes from the bottom when arranging them on the PCR tube rack. A clean pair of forceps can be useful for this. The tube rack should be clean and wiped with a DNA decontamination reagent before you start.

• Do not splash or spray PCR samples.

• Change gloves frequently especially if contamination is suspected.

• Keep reactions and components capped whenever possible.

• Premix reagents before aliquoting into individual reactions to reduce the number of pipetting steps

• Minimize opening of PCR tubes post-amplification. Gel electrophoresis of PCR products should be carried in a separate lab. Real-time PCR systems eliminate the need to open reaction tubes for analysis.

• Once you have finished aliquoting your PCR master mix into the PCR tubes the water should be added to the negative template control tube in the pre-PCR clean room. All reaction tubes should then be closed and taken into another lab where the DNA template is added to the rest of the PCR tubes.