Molecular biology is a tool. Successful working of this tool depends on many factors (sometimes beyond the control of a molecular biologist). Therefore, to increase the chances of success (and to avoid silly mistakes unknowingly), I shared some useful molecular biology related tools and resources in my earlier post. Have a look.
I experienced that, the PCR related tips which I am going to share in this InfoMention are not new and exhaustive to many of us. But sadly, only few of us would try these at the time of desperation like I portrayed above. Simple ignorance can be one of the reasons for this. If even after following these tips, you are unable to get the required PCR product then there is a volume of literature available both online and in library. Feel free to refer them too.As I don’t know the basic skill sets of my audience for this post, so to give a fair chance even to a new molecular biologist I am starting from the very beginning and very basics. Please skip those sections which you already know.
I recommend that you should always order primers as desalted and lyophilized dry state. You can choose 0.05umol as synthesis scale (although 0.025umol may also work, but remains risky). Centrifuge briefly these dry tubes before opening the cap. This ensures that the entire dried pellet remains at the bottom of the tube. Always use tips with filter cap. You may contaminate the primer stock by using tips without filter using the same old pipette from the lab. This can pose many problems in future.
Add the equivalent volume of “filtered Tris.HCl (10mM, pH 8.5)”. What I mean by this is, if your primer tubes read as 42nmol then add 42ul of Tris.HCl. If it reads as 68nmol then add 68ul of Tris.HCl. By doing this you will have a 1mM stock of individual primers (Fwd and Rev Primers reconstituted in their respective tubes). This makes further calculations much easier.
Tip 1: As we are going to use both Fwd and Rev primer together in each PCR reaction then why to add them separately each time for each PCR reaction. To this, I can suggest that you make a “primer-mix”. Do this separately from the original primer tubes. In my experiments, I use to make “primer-mix” at a concentration of 5uM by dissolving 1ul of Fwd and Rev primers (from above 1mM stock) in 198ul of Tris.HCl or Water.
Most researchers buy already prepared dNTP mix. But I recommend that you should make your own dNTP mix using 100mM dATP, dGTP, dCTP and dTTP (for this you need to buy these individually as 100mM). To make 5mM dNTPmix, just add 5ul of each 100mM dNTPs (individual) into 80ul of water and mix them well by brief vortexing. Commercial dNTP mix sometimes may not have exactly the equal concentrations of each dNTPs which may affect very sensitive PCRs. Remember; we are talking here about PCR troubleshooting!
Mostly (whatever be the reason), I have seen that people do PCR reactions without DMSO, and at times they see no desired bands. I think, we can overcome this by the use of DMSO gradient. By this, I mean use of DMSO at different amounts in each PCR reaction. I often prepare 100ul of PCR reaction mixture suitable for 4 PCR reactions with 25ul in each tube. Then I use to add different amount of DMSO e.g. 0, 0.5ul, 1ul and 2ul DMSO in my 4 tubes of PCR reaction. This way, all 4 PCR tubes will have exactly the same PCR reaction constituents except the concentration of DMSO. After working with many clones and different vector systems, I found that using DMSO gradient (sometimes from 0 to 4ul in each PCR reaction tube) not only helped to save my time but also gave cleaner bands (in contrast to smear, false amplification and primer dimer bands etc). One may argue here that this is simply the waste of PCR chemicals because we may only need one PCR tube. To this, I would like to say that in Research, Time is money! If you can save your later time now, then you are a winner otherwise you will waste both time and money (chemicals) later on in unnecessary troubleshooting.
Always (without fail) do control PCR (i.e. without template DNA) along with any experimental one using the same set of DMSO gradient. If both experimental and control PCRs show the same molecular weight band of your interest please don’t be tempted to still use it. Trust me, it’s of no use. However, if the control PCR has other bands (not closely matching with the band of interest in your experimental PCR reaction) you may still use the experimental PCR but do that with prudence.
Tip 2: DMSO is used as a PCR enhancing agent which prevents the formation of secondary structures of either primers or template. Betaine is also used as a PCR enhancing agent but from my experience use of DMSO is cheaper and hassle free.
I use pfu-turbo DNA polymerase for my PCRs. I know this is costly, but it’s worth using it. I recommend using any DNA polymerase which does not add any extra overhangs (e.g. avoid Taq polymerase). Sometimes, one may get good amplification only by changing the DNA polymerase (although no one would tell you this, but I have personally experienced it). Just think if one high fidelity DNA polymerase would suite all PCR types then these companies would not sell ranges of DNA polymerases.
Generally, the buffer which comes with these DNA polymerases works well in most conditions, so you can use it. But, I found KLA buffer works best (sometimes better than the supplied buffer). The final filtered KLA buffer at -20C is stable for more than a year or so. You can prepare 10X KLA buffer as follows:
500mM : Tris-Cl (pH 9.2)
160mM : Ammonium sulphate
25mM : MgCl2
1% : Tween 20
Many people prepare 25ul of one reaction mixture. If things worked well for you then no need to change anything. But, if you are going to try something new or things which worked in the past suddenly stopped working you may try my method.
Here, I suggest preparing 100ul of PCR reaction mixture and then dispensing 25ul in 4 tubes having DMSO gradient (0ul, 0.5ul, 1ul, 2ul). If you do not see bands with these 4 DMSO amounts, then you can even go higher with DMSO (e.g. 2.5ul, 3ul, 3.5ul and 4ul). I have not gone beyond 4ul DMSO. I used 4ul DMSO to PCR amplify GC rich template. This you can easily find out using pDRAW software. Have a look.
To prepare a 100ul PCR reaction mix, I use to setup the following:
1. Water: Xul to make total volume as 100ul (without DMSO)
2. 10X KLA: 10ul (as MgCl2 is already in the KLA, so I am not disturbing it). If you use default buffer, then you may need to titrate magnesium concentration.
3. 5uM Primer-Mix: 4ul
4. 5mM dNTP mix: 4ul
5. DNA template: 200-500ng (so that each tube will have at least 50ng or more).
6. Pfu turbo: 0.4ul (strangely, unlike many other DNA polymerase, 0.1ul of pfu turbo is more than sufficient in each tube for normal PCR).
Mix well this above PCR reaction mix and pipette out 25ul each into 4 PCR tubes already containing (0, 0.5, 1, and 2ul of DMSO). Keep all tubes (reaction mix and individual tubes) on ice during all steps.
Tip 3: Here, you can sometimes differ the amount of primer-mix and DNA template depending on your results. Meaning take less primer mix (e.g. 2ul instead of 4ul) if you are getting too many non specific bands.
Tip 4: Another option is to use a fresh DNA template or you may linearize the DNA template for clean specific bands.
Unfortunately, this is listed in the troubleshooting section of most of the PCR vendor’s guide and not in the prime step section. I use this “touch-down” on routine basis for all my PCRs. Touch-down is nothing but some extra cycles which the machine runs before the actual 30-32 cycles. In the touch-down step, we program the annealing temperature to go down by the mentioned temperature (e.g. 1 or 2 degree on each cycle) for the mentioned number of cycles (e.g. generally 5 to 10 number of cycles) before it actually starts the number of main cycles. This ensures the proper annealing and initial extension of the primers.
If you think annealing of primers could have been a problem, then increasing the annealing time from say 30s to 1.4min sometime helps.
Finally, during the last extension, the default time of extension at 72C is for 5min. Just increase the time from 5min to 8min and see the change.
When your PCR is finished, make sure that the Lid temperature also falls, so program the machine accordingly.
This is very interesting. Not many labs have an opportunity to have two different brands of PCR machine. I was lucky to get this experience too. I had some problem in PCR amplification. As I had my program written on one machine so I used that machine always. But in one of my tricky PCR (even after trying all the above mentioned steps) I was always getting false amplification. At that time, I was using only one PCR machine for all my troubleshooting. One day that machine was not available in the time slot I wanted. So I had to write the same program on the other machine. And to my surprise, I got my PCR amplification the first time I did it.
Afterwards, for my peace of mind, I ran the same PCR in parallel on both machines to repeat the result. I got PCR amplification in one machine again and not in the other. After investigating it in depth, I found that the machines differ in the “temperature ramping speed” and perhaps responsible for that different result. Generally (for some sophisticated newer models), you do have an option of playing with the ramping speed too. I suggest to try this when you are left with no other option.
At the closing remark, I found Steve Albini lines very appropriate:
By the way, want to know all about cloning a gene and its troubleshooting, then watch out for my next post
Until then, wish you a successful PCR.