It is possible that you are detecting primer dimers in your NTC. Here is a simple Thermo reference. https://www.thermofisher.com/us/en/home/life-science/pcr/real-time-pcr/qpcr-education/real-time-pcr-troubleshooting-tool/gene-expression-quantitation-troubleshooting/amplification-no-template-control.html

 I have attached one of the results I have been having. The expected PCR band is 150bp and you can see the same band in my reverse transcriptase negative and my NTC for the PCR reaction. I don't think these are primer dimers because I would have observed these before contamination became an issue. Also the primers are used by my other colleagues routinely.

In my lab some grad students always were using UV to decontaminate pipettors. And I heard many times dna contamination was an issue for them in qPCR. Try to clean your reagents, pipettes etc. It looks like you have cDNA cont.

Good luck I hope it helps... 

There might be contamination in pipette tips so autoclave them before use (make sure they are autoclavable). To prevent contamination from the pipette itself, use plugged pipette tips.

Another possibility may be a contamination in your plate, make sure these are sterile too.

Hope this help!

You said the primers are used by your colleagues, do they run the same PCR and get no signal in RT-ve and NTC? In any case, if you share the primers, that might be a reason for concern, I would just order a new batch of primers, and take an aliquot inside a class I or class II hood, then work only off that aliquot.

Have you worked with a plasmid containing the cDNA you're testing for? If yes, did you handle the tube containing the plasmid in the same area as you set up the PCR? If yes, micro-aerosols from the tube could have contaminated a lot of things in your environment, including things like labcoat and tube racks, which would be bad news (happened to me once, I had signal in water NTC every time since I had used a plasmid with the target gene).

Try to separate the PCR setup stage and the gel running stage as much as you can: separate set of pipettes and tips, separate area of the lab (or ideally a separate lab), separate labcoats.

And as people already said: use filter tips and UV-treat your pipettes.

Good luck and let us know of your progress!

In February we moved premises and though I don't work with plasmids or do any cloning my colleagues either side of my bench definitely do but I don't think they would be cloning this fragment. We just PCR it to check cDNA quality and that the negative controls have worked. The 100uM primer stocks are communal yes but we keep our own 10uM primer aliquots. Yesterday morning I UV radiated my pipettes for 50 mins prior to making the master mix and this seemed to remove the positive band in my NTC. Thus, I carried out  PCR with exactly the same reagents for my cDNA samples, RT neg and NTC and the band reappeared to my annoyance.

I know what you're going throuhg. But I like how enthusiastically you names the file with clean NTC ;-)

Here are some more things to double check. Add the master mix to the PCR tubes/well first, and then you add templates. Once a tube with mastermix and template is closed, you don't open it again.

This one is contentious: add your NTC and RT-ve templates first, close the tubes, then proceed with the test samples. The reason this is a good idea is because you will be more likely to avoid your NTC being contaminated by aerosols from other samples. This will be more likely to give you a clean NTC. The reason this is a bad approach is because there is something causing cross contamination between tubes/wells, you do want to know it and a signal in the NTC might be telling you that. In other words, if you use this method to prevent your NTC contamination but you actually do have an underlying aerosol problem, you won't know it and some of your apparently positive samples might just be positive as a result of aerosol contamination. This is why some recommend that NTC should be the last template to be added.

Three more questions:

1. How do you mix your samples: do you vortex them or pipette up and down?

2. When your colleagues use these primers, are their water and RT-ve controls clean?

3. Do you run your PCR in PCR tubes or plates?

Hi Andrzej. Thank you for helping me I really appreciate it. To answer your questions I vortex my PCR tubes prior to heat cycling and my colleague ran this PCR today for cDNA, genomic DNA , NTC and my RT negative sample and got perfect results. Unfortunately though the RT negative came back positive so this suggests that my cDNA samples I made a few weeks ago now also carry the same contaminant. I run all my PCR in tubes rather than plates and I have attached my colleague's PCR for comparison. Apologies for the glare.

Then my last recommendation is a general clean-up: change your lab coat, order fresh reagents (including RT reagents and oligo dT primer), UV-sterilise whatever you can, and what you can't - wipe diligently with mild bleach, e.g. DNA away. New tip boxes, new batch of tubes, etc. Set up a new experiment from which you will isolate new RNA using a new RNA isolation kit (after all, it's your RNA that might be contaminated).

I know this might seem like a waste of your current consumables, but in the end you'll end up wasting more resources trying to figure out where the contamination comes from exactly. Try to do your RT and PCR set-up in a class I or class II hood if there is one available, as this will minimise the risk of contamination. Good luck!

To echo what everyone has said, throw away ALL OF YOUR REAGENTS (primers, buffers, water, enzymes, etc.). Don't share reagents/tips/pipets with anyone. Have a dedicated pipet to load your gels and don't use it for anything else. Contamination is a miserable problem to have. But the good news is that you only care about fixing the problem, you don't need to discover where it came from. Good luck!