DMSO is a PCR additive that helps denature the template DNA. We use it in PCR and sequencing for GC rich templates. We have tested percent addition and found 5 percent is generally optimal for most amplifications. I would not recommend less than 3 percent or more than 5 percent. Adding DMSO should not have a negative effect on amplification even if the template is not GC rich.

I would have to agree with some of the other comments. The Tm values of the primers should be relatively close, maybe 4 degrees different. You could also check the sequences of the forward and reverse primer. One note of caution. Primer dimer does not necessarily have to occur between the forward and reverse primer. It could also occur when the forward primer has complimentary regions with itself and the same for the reverse primer.

Firstly vary your annealing temperatures accordingly (gradient) and see.

If still problem persists, add forward primer for say 5 cycles first and then add your reverse primer post 5 cycles. This ensures you have some initial product from your forward first and thus, the probability of increasing your target amplicon after reverse addition, hence, avoiding primer dimers

hey first thumb rule is your Tm of F&R primer should no vary more then 5 oC.

if u r not getting primer dimer means the following things might be possible

1. Insufficient DNA template

2. too low annealing temp.

3.complementary binding sites between primers

4.contamination of reagents etc.

remedy

1. try increasing DNA template or Tm

2. redesign your primer. you can use this tool for designing primer

http://eu.idtdna.com/analyzer/Applications/OligoAnalyzer/default.aspx

i always use that..

y can even try adding DMSO

Dear Biswajit

I agree with mohamad imran that you try a gradient PCR 1st with same conc of MgCl2 at which you got some amplification. If you get primer dimer even after that, then try the following, which is from my personal experience:

Mix your primers and water (only the amount you need for current PCR reaction), heat at 65 deg for 5 min and then snap-cool in ice. now add rest of your PCR reaction components and perform the PCR.

Thanks and all the best

you are taking more primer concentration then required or you are taking less template DNA. if you want to do sequencing then you can cut your product from gel and purify it prior sequencing.

You can check the sequence of your primers for the probability of dimer formation. There are several programs on the internet (one of them is this one: http://yellow.nist.gov:8444/dnaAnalysis/primerToolsPage.do). If you have a certain degree of flexibility with your primers, just design new ones which aren't prone to dimer formation....

Hi Navjyoti, your method is quite novel to me. can you explain the function of this reaction in PCR programme.

try oligoanalyzer.. than use gradient PCR for right annealing temperature... maintain the same concentration of ions at which u r getting ur product also besides the dimer....

Hi navjyoti,

Even I wish to know the role of snap-cooling primers and water mixture before proceeding to pcr

Here are my suggestions: Try to reduce five cycles of PCR reactions. Try to reduce primers or increase DNA. Increase Tm.

Thanks Ankur ...What will be the probable % of DMSO .....Is it ok with DMSO for sequencing...

What i would suggest is that : 1) When you are designing primers then both the primers should have anealing tem almost equal or 1-2 degree difference. If both the primers having differece in Tm then anealing tm of lower should be used for PCR. If dimer is comming then use tm of higher one or 2-3 less as in your case difference is of 9 degree.

2) In your case long PCR can be tried with tm starting from higher to lower one.

3) If still you are getting the same result then you can do PCR of bulk sample (50ul) and load all the amount on Gel, cut the band , elute out, check on gel again and then send for sequencing. (this is the quick and easy method).

good luck

1. Increase anneal temp.

2. Add primers to master mix one at a time, all steps on ice.

3. Use a hot start Taq.

4. Run a Mg gradient.

5. Redesign primers.

DMSO is a PCR additive that helps denature the template DNA. We use it in PCR and sequencing for GC rich templates. We have tested percent addition and found 5 percent is generally optimal for most amplifications. I would not recommend less than 3 percent or more than 5 percent. Adding DMSO should not have a negative effect on amplification even if the template is not GC rich.

I would have to agree with some of the other comments. The Tm values of the primers should be relatively close, maybe 4 degrees different. You could also check the sequences of the forward and reverse primer. One note of caution. Primer dimer does not necessarily have to occur between the forward and reverse primer. It could also occur when the forward primer has complimentary regions with itself and the same for the reverse primer.

I noticed that you used very high conc. of MgCl2, I normally used 2 - 3 mM MgCl2 (final conc.) in PCR reaction. You also mentioned about universal primers, do these primers have degenerate nucleotides in them? If so, it will be very difficult to avoid primer dimer, but you can reduce the primer dimer formation by reducing primers conc. (not sure how much you are using) and conc. of MgCl2 in addition to try gradient PCR as suggested.

check with primer design.if its right, go with different concentrations of Primer, Taq, dNTPs aswell....keeping a standard mgcl concentration.or you can manipulate annealing temperature.ususally primer dimers are reduced with increase in annealing temperature.but you can aswell check gradient pcr (keeping all other conditions constant)..usually between 52 - 65 degrees.

if all this doesnt work, try reducing template DNA concentration. goodluck.

If the PCR product is over 300 bases you could just use Ampure beads to remove the primer dimer prior to sequencing, if you do the conditions right then the beads won't bind fragments less than 300 bases long. That might be more efficient than spending time trying to optimize the PCR, unless you plan to sequence hundreds or thousands of samples that might make it worth the time to optimize the PCR.

If you prefer you can also perform a PEG precipitation...but your template must be over 250-300 bp. Here it is the protocol. It is easy and fast and to eliminate your dimer problem without optimized the PCR (as proposed also by Tim Smith):

Stock Solutions:

1) 20% PEG, 2.5 M NaCl - Note: PEG takes 20+ min to go into solution when you make the initial solution. For 50mL, mix the following in a 50 mL conical:

10.0 g PolyEthylene Glycol 8000 (MW = 6000 - 8000 is fine);

7.3 g NaCl;

dH2O up to 45 mL - shake & let PEG go into solution; I put it into a rocking incubator set to 37°C after everything is in solution, fill with dH2O up to 50 mL.

2) 80% EtOH in a 50mL conical or something similar.

3) TLE = 10 mM Tris, 0.1 mM EDTA

or distilled sterile water (I prefer...)

This protocol assumes a 50 μL PCR reaction and use of 0.2 mL thin walled tubes. If you have a different volume of PCR, then scale everything using the following proportions. If you use 0.5 mL tubes for PCR, then you don't need to use any new tubes.

A) Run out 5 μL of PCR onto an agarose gel to ensure the PCR reaction worked.

B) Add 50 μL of PEG to a 0.5 mL tube. Transfer the remainder of the PCR to the tube with PEG and mix

by pipetting up & down.

C) Let the the PCR + PEG incubate at 37°C for 15 min.

D) Place the 80% EtOH on ice.

E) Centrifuge PCR + PEG at high speed (~15,000 x g) for 15 min. (at room temp.).

F) Using a P200 pipetter, pull off the supernatant & discard it.

G) Add 125 μL of cold 80% EtOH to the tube. If you shoot the EtOH into the bottom of the tube, you must spin for two minutes. If you place the EtOH onto the side of the tube, you can just let the tube sit for one minute. Using a P200 pipetter, pull off the supernatant & discard it.

H) Repeat step G.

I) Dry off the EtOH in a centrivap for 5-10 min, low heat, no vacuum. There should be no trace (visible

or by smell) of EtOH when done.

J) Dissolve the PCR product in 25 μL of TLE or gdw. Pipette up & down several times to ensure the DNA has gone into solution. If you can let it sit for several minutes, that is also helpful.

K) Run out 2-4 μL onto an agarose gel for 10 min. to roughly quantify recovery.

I hope it is clear. :-)

Thanks a looot @ Doug Bintzler Sir....I will try with 5% DMSO. In PCR mix (25 μL) can I use 4-5 μL of DMSO and after preparation OF 5% DMSO, I have to autoclave the DMSO solution ?

Thanks Baochuan Lin Sir.....I will go for a gradient MgCl2 conc.

First, the Tm of your primer are really different, if possible I would redesign primer with PrimerBlast (ncbi), if not having the possible to change primer sequence you might change or add a tail in order to modify primer Tm, additionally the primer conc. should be in a range of 0.1 to 1 uM, and MgCl concentration are indeed very high, so try reducing them to 1 upto max. 4 mM. Another pont of primer dimers could be a to low Tm or a to active Taq- Polymerase, so maybe you could also reduce the conc. of your enzyme. Maybe touch-down PCR could also help.

There are many good suggestions above.

Have you checked the complimentarity of your primers? Perhaps trying to understand why you have primer-dimer formation will help with the strategy to avoid it.

Often a temperature gradient (if you have an available thermocycler with that option) using a single PCR mix gives good results. A two temperature PCR program may also be a good option, try a joint extension and annealing step with a temperature around 66ºC. But sometimes, if it is feasible, it´s best to chose another set of primers.

The Tm for Fwd and Rev primers should be closer (preferably less than 2ºC); reduce primer concentration and adding up to 10% DMSO may help.

well, i have tried DMSO for my previous such problem...it didnt work much though! infact, there was no amplification atall...check with high annealing temperature...with minimal quantity of template DNA and Mgcl2.

I don't agree with Tin-Chun. Fwd and rev primer less than 2°C?! Then the most PCR-applications won't work.

well, i agree to nandine....usually we manipulate Tm +5 or -5 degrees or even more/lesser sometimes.

It seems there is a big difference in the melting T of the two primers. Try to perform optimisation by varying the Mg concentration, primer and DNA concentration and temperature. Sometimes hot start polimerase helps!

I have tried with 2mM Mg+ and 2pM primer conc. with 5% DMSO. I got some smear but no band. I didn't get the result (bands) that i previously got. But, got reduced primer dimer with low Mg+. Should I go beyond 60 degree and can I go with the lower Tm or higher one of the above mentioned primers( below/ above)?

The lower your PCR annealing temperature is, less specific it`ll be. If you go much lower the unespecific problems will continue. If your PCR annealing time be too long, you may increase the likelihood of unspecific amplification products.

Check with your primer concentration. If you use less primer concentration, It will helpful for you to overcome from this issue and also try hot start PCR

Well in general i think the first step will be to optomize the concentration of the primers so that will be just enough to anneal with your template DNA.I will suggest to try a range of three primer concentration (with differences of 0.5 micrmol) and keeping other conditions as they are. One of them should show less primer drimer and could be used as indicator to go below that concentartion for optomoize your experiment. If that didn't work then you migght need to change one of your primer pair

-Try hot-start PCR (with a hot-start enzyme or in putting your tubes in the thermocycler when it has reached 94 °C)

-Design new primers with closer Tm; use lower concentration of primers and of MgCl2 (we usually use 1,5mM as final concentration !).

Sorry, you can eliminate your dimer after the PCR whit a simple purification with column kit or using a simple PEG precipitation.

I attach a protocol in a similar question. Now I am at home and I must serch....waiting..

-simply do two things first try out different primer conc...if you are using high conc of primer than u will get primer dimers, put primer conc. gradient reducing your primer conc upto 1-2 pmol/ul.

Second reduce your pcr cycles.

try these ...

well, u got no bands possibly becoz of DMSO, which is a strong denaturant. anyways, try 2mM mgcl, 2pm primer conc, and high annealing temp....why dont u go for gradient between 55-65 degrees temp.and do not add DMSO this time. see if this works.

Look my brother, the primer should not be so lengthy that u will gel easily the primer dimer, u have first check wheather you are finding any amplicons, and if u r getting some u should test 67 degrees as the annealing temp, i hope it will work, as for my self i trouble shooted the same problem...rest can also depend.

You may have Dimers because your primer sequences have complementary sequences. You may anticipate this kind of situation already when making the choice of the best primer pair. Bioinformatic tools will tell you which pair as lower risk of dimers or which one has dimers that may be easily split during standard PCR. I would suggest you first make simulation for your Primer pairs using Vector NTI (licensed Bioinformatic tool). This tool will foresee the number, type and position of Dimers that may result from the use of each primers pair. You will also get the amount energy required to split each dimer. The Primer pair with Dimers requiring energy around 0 is the most efficient.

Just looking at your melting temps, the primer pair you are using appears to be far less than ideal. When designing primers you want the melting temperatures to be very close to each other. I always used the old school way of A or T contributes 2 degrees and G or C contributes 4 degrees to the Tm value and generally design primers that are 21mers with Tms of around 52 - 56 degrees. More modern analyses methods take into account salt and Mg concentration in your master mix, formation of hairpin loops etc. based on submitted sequences and fancy algorithms but I have rarely had failed reactions with the old school method of calculation based on visual inspection of the DNA sequence in need of amplifying. Certainly the fact you have a difference of 11 degrees between your primer pairs is a problem and the formation of primer dimers is likely caused by significant complementary pairing of your primers. You should assemble your PCR reactions on ice. In addition, you could also use a DNA polymerase that is only activated in the heating step to improve non-specific amplification. As far as your primers are concerned, I would cut back on the nucleotides of your reverse primer so that it's closer to the Tm of your forward primer (54 - 56C). As a general rule, an annealing step of 5 degrees below the Tm value, which in this case is 50C, should work. And remember: the success of your PCR is directly proportional to the robustness of your primers.

Low temperature mis priming can be avoided by initiating PCR at an elevated temperature, a process usually reffered to as Hot start PCR.This may be performed by introducing a critical reaction component,such as polymerase after the temperature of the sample has been raised above the desired annealing temperature e.g.60 degree centigrade.

Primer dimers are very common when using universal primers. But i dont see the problem you are facing... If the purpose is sequencing then i would suggest you to clone the gel eluted product into any TA cloning vector. If vector availability is a crunch, then it would be better if you perform the PCR reaction at higher annealing temperature for longer time (may be for 1min 30sec). Primer dimers can also be reduced to some extent if you perform the PCR using the gel eluted PCR product as template also. All the best...

After primer designing, I check the probability of dimer primer formation in http://eu.idtdna.com/analyzer/Applications/OligoAnalyzer/

In this web site, you could calculate kcal/mole of slef dimer, hetero dimer, hairpin formation of one sequence of primer or pair set of primers.

What are the annealing temperature you have used for your study. Small sugg. its better to keep TM of both the primers close to each other and then use high annealing temp to avoid the PD issues and also to make the PCR more specific

Do you necessarily have to use those primers? If you cannot design an alternative pair of primers, try the suggestions on the attached paper. Sometimes I use BSA instead of DMSO.

I usually design primers with Primer3 (v.0.4.0) and check them with Oligocalculator (http://www.basic.northwestern.edu/biotools/OligoCalc.html), besides of the advice of Ms. Samaneh Tousi. Just in case that you can design new pairs.

Article Multiplex PCR: Critical Parameters and Step-by-Step Protocol

Differeence between two Tm value is very high. Thus you try for a touch down PCR sarting 65-54 °C.

Look for primer sequences and consider primer dimer's lower thermodynamic stability. Keep your annealing temperature much higher than the TM of primer dimer. That means one have to be careful while designing primer.

If you are doing for cloning studies and using restriction enzyme seq at end try changing them to minimize the Tm gap. Like using AT rich at reverse primer or GC rich at forward primers.

OR, try redesigning the prime little bit upstream of the sequence. I usually use OligoAnalyzer for designing primer. here is the link..

http://eu.idtdna.com/analyzer/Applications/OligoAnalyzer/default.aspx

one more thing.. ideally tm should not vary more than 3 to 4 oC

Mg concentrations are 10 times to high.

You increase annealing temperature

Determine the lowest primer concentration that works for you. Usually, together with annealing temp adjustments, this will get rid of your problem. But We find that by simply lowering the concentration of your primers in the PCR, you get rid of dimers.

If you must use this particular pair of primers, try setting up a titration experiment by changing the concentrations of the upper and lower primers, for example the ratio of Upper primer:Lower primer is 1:3, 1:2, 1:1, 2:1, 3:1. Also try experimenting with PCR cycling conditions (annealing and extension temperatures and time; and perhaps increasing the number of cycles) as well as the concentration of MgCl2.

Primers are much cheaper than your time, I would redesign the primers and see if that helps. There are a lot of free tools that will assist in primer design and still give you the option to pick between the suggested options.. IDT (integrated DNA Technologies) has a great tool on their website that is free and provides different choices of primer pairs. They also have a section where you can specify exactly where within your target sequence you would like your primers to bind so you can target regions within a gene. You can adjust your Mg++ concentration and other PCR conditions to aid in primer design. It will also evaluate the probability for primer dimers.