Hi all,
I am currently troubled with my PCR assays because what used to be a successful amplification and visualization of my amplicons has suddenly stopped working. Here is the following protocol I have used for all my PCR assays. The one I show below targets the 16S gene of Bacteroidales spp, with an expected amplicon size of 170 bp. The DNA template solution I have used is extracted from mice feces. Here is the following master mix and thermal cycling protocols:
Master Mix (per 50 uL):
- 5 uL of 10X Thermopol Buffer
- 1 uL of 10 mM dNTPs
- 1 uL of 10uM Forward Primer
- 1 uL of 10 uM Reverse Primer
- 1 uL of 200 U/uL Taq
- 2 uL of DNA template
- ddH2O to 50 uL
Thermal Cycling Conditions:
- 95oC; 5 min
- 95oC; 30 sec
- 52oC; 30 sec
- 72oC: 1 min
- Repeat Steps 2-4 35 times.
- 72oC 6:00 min
I have attached 2 files, each file name being the date I conducted the assay. 011214.tif is an image of a successful gradient PCR I conducted with the primer pair. Assays previous to this one were successful in amplifying the desired target. UniBac010615 is a normal PCR assay I conducted today. The only difference is that I halved the reaction volume to 25 uL. The volumes were changed to reflect the ratios, the only exceptions being that I still added 2 uL of DNA template and 0.25 uL of the Taq solution to each reaction mix.
I have also obtained a different stock of Thermopol Buffer because I ran out of the buffer. Other possible causes I hypothesized are the excess Taq enzyme, impure primer solution, and impure water.
If you guys have any other suggestions, I would welcome them.
Thanks!
See ur taq conc. It seems more.
My suggestion is to use new taq with buffer and MgCl2. Make new primer working solution and dNTPs. See what happens. Otherwise u have see them one by one.
Hi Poul
You reduced the Taq concentration by a factor of 2. That could reduce the total amount of amplified DNA possible. In addition, it is conceivable that the DNA template is contaminated by something that inhibits PCR; by lowering the reaction volume, you effectively double the concentration of those inhibitors.
The PCR DNA migrates very close to the dye. In the first gel, the DNA migrates below the dye whereas in the second gel, it migrates right in the middle. This is likely caused by small difference in agarose concentration or running time but can affect the visualization of your DNA quite a lot.
I suggest you repeat the PCR and maybe set up a few control experiments (lower template concentration, different primer pair etc.)
Best
Jesper
Hi Paul,
I think the products you are showing are at the size of 170-200 bp which is what you expect, however to make sure I think you should run a positive control such as GAPDH to make sure that your master mix anDNA template are working well. You need also additional tube with no DNA template to show that your dNTPs, water and others are clean and show no contamination.
To avoid changes in all the PCR reactives is better to use the master mix 2X ready to use, just add the template 10-30 ng and primers at 10 uM, there are from different brands and it will be easy for you, it works in 6 uL volumen or 12, 25. I use a lot PROMEGA brand, green master mix.
Hi Paul,
What was the concentration of the DNA you used? It could be that you don't have a sufficient amount of DNA to start the PCR with.
Have you used a positive control? In this way you can see if something went wrong during the reaction (for example you forgot to include your taq polymerase or like previously suggested, you need to use new taq polymerase).
I would definitely change the Taq as an initial easy fix, it doesn't tend to appreciate being left out at room temperature for any length of time.
May be the wind was blowing the wrong direction that day? I hate when this happens. However, whenever you change the total volume of reaction, unless you ratio it down to all components could lead to unsuccessful reaction. DNA template and primer ratio is very critical in PCR amplification. Keeping the same amount for half of the reaction volume may have caused it? so I would try with same ratio reduction and keep one tube of 50ul reaction as your positive control and would also always include negative control and try again. This will help you answer the question if ratio was responsible.
Dear Paul,
In 50 ul of the reaction the dNTPs you are using seems to be very low than the required. Also you used 1 ul of Taq (200U/ul) which means you wasted 200 reaction only for setting a single reaction which is a great loss. The ratio is very important. the primers, dNTPS, etc. if you are setting a PCR of 50ul then 1ul of 10uM primers may be too low also same case with the dNTPS where you used 1ul of 10mM dNTPs for 50ul reaction which is againg very low. you should have added 5ul of 10mM dNTPs in 50ul reaction.
One thing you can do is that freshly make aliquots of the components like primers, Taq, dNTPS, etc from a new stock then run a small scale PCR of 10-20ul where you can added 1U of enzyme, 1ul of 10-20uM (pM/ul)of primers, fresh dNTPs and also you can add some additives like MgCl2, 1% DMSO, BSA . the DNA should be at least 30-50ng/ul and run few reaction.
All the best
Hi
All recommendations are nice. Another possible error is using old thermal cycler machine that can not detect the reagent level automatically. If the problem did not solved with mentioned keys check the performance of your PCR machine.
Best
Thanks for all the suggestions everyone,
I managed to solve the problem with the UniBac assay.
However, I'm experiencing the same problems when I use 16S primers (target all bacterial sequences for next-gen sequencing). I'll follow all the suggestions and keep you all updated.
- Badges
- Science method