Hi

one reason for diluting out DNA sample prior to PCR is to negate the effect of inhibitors: Sometimes if you use too much DNA, e.g. > 200ng of genomic A for example for PCR amplification you gain nothing in terms of extra copies of your DNA contributing to efficient PCR but in effect by adding more you over load with inhibitors; in particular for most DNA preps salts: Generally speaking inhibitors tend to behave according to zero order kinetics; that is below a certain threshold they fail to inhibit but above this level they prohibit amplification. Thus you can have a situation where you fail to obtain bands during PCR at a certain concentration, e.g. 100ng of genomic DNA but if you try 20ng the PCR will work

In addition, by limiting the amount of template in certain instances you limit non specific binding of primers and extra bands in PCR. In this case too much DNA can lead to a strong specific bands but extra non specific bands whereas less template just results in the one specific band 

Huge concentrations of DNA do not let reaction to flow normally, e.g. they inhibit reaction. For example, on the agarose gel you may find big shining plums (so called smears) of non-amplified DNA scattered along the row.

Hi

one reason for diluting out DNA sample prior to PCR is to negate the effect of inhibitors: Sometimes if you use too much DNA, e.g. > 200ng of genomic A for example for PCR amplification you gain nothing in terms of extra copies of your DNA contributing to efficient PCR but in effect by adding more you over load with inhibitors; in particular for most DNA preps salts: Generally speaking inhibitors tend to behave according to zero order kinetics; that is below a certain threshold they fail to inhibit but above this level they prohibit amplification. Thus you can have a situation where you fail to obtain bands during PCR at a certain concentration, e.g. 100ng of genomic DNA but if you try 20ng the PCR will work

In addition, by limiting the amount of template in certain instances you limit non specific binding of primers and extra bands in PCR. In this case too much DNA can lead to a strong specific bands but extra non specific bands whereas less template just results in the one specific band 

Dear Laurence Hall and Artem  Skrypnyk

many thanks for your nice and quick response. I have been using the diluted DNA samples with the concentration of about 100ng/micro litter and the result was not that much good. I will try to reduce it to the level 20ng and try it,   hope it will work and let you two know the result.

Many thanks again

Dear Tesfaye

Have you tried validating your DNA with primers to a housekeeping gene like b actin or gapDH that have worked in prior DNA before trying with your GOI primers; In effect that constitutes a positive control. Indeed have these primers worked before on other DNA samples which in effect validates the primers and therefore implies that the problem is your DNA sample and not your actual primers.

When I perform PCR I tend to do 3 basic things (inter alia):

1. Make sure the 260/280 ratio is 1.7-2.0 (implying low protein contaminants) and more importantly verify that my 260/230 ratio is > 1.0 and preferably > 1.5 implying low salt contamination. In many instances this is actually more important than the 260/280 ratio which most people tend to fixate on

2. That said, with well designed primers, I routinely genotype from crude genomic cell lysates, not pure (purified) genomic DNA where sat and cell proteins are removed and that works perfectly well with PCR conditions that are optimised in terms of annealing temp and primer design: Just to cover the concept of good primer design IO have attached a primer design doc that you might find useful. In particular, with new primers, regardless of cDNA amount and quality PCR can fail because

A. Your primers bind to each other (primer dimers) limiting the effective concentration available for PCR. This tends to be because primers end with 2 or more G/C residues which leads to stable steady state primer dimers

B. Bind to themselves either through complementary G/C bases at the 5' and 3' end or internally causing hairpin loops. Thus, this tends to be an issue with GC rich target sequences where primers are consequently GC rich

C. Terminate primers in multiple A/T residues which leads to weak priming and poor efficiency PCR

Thus you design primers with a certain (optimal) base composition and screen for such structures. See attached

 D. Design primers with identical Tms or Tms +/-2C and try 20-100ng of genomic DNA with an annealing temp of Tm-2C: In particular therefore most optimal primers will be 18-21bp in length have an average GC content of about 50%, leading to Tms of 60C to 65C which therefore lend themselves to annealing temp of 58C and 63C respectively (by way of example)

E. Remember also to check your target sequence in genomic DNA you are attempting to amplify from: if the GC content is above 60% or the region you are attempting to amplify contains runs of more than 4-5 GC residues then PCR efficiency without additives like betaine or DMSO or use of prolonged high temp denaturing conditions, e.g. 96-98C can result in inefficient PCR. Let me know if you would like me to provide further details

I have stressed primer design and selection of good target site because with optimal primers as described in the document; optimal PCR including the use of Tm-2 for annealing and the use of generally 20-50ng of gDNA PCR in my experience works consistently well even with crude genomic lysates

Finally, if you have primers that are optimally designed and moreover have worked with other genomic DNA samples, implying the issue really is a sample issue then check as mentioned the 260/230 ratio. If you find it is less than 1.0 then you can remove impurities (salts) and hopefully enable PCR to finally work by:

1. precipitating the gDNA and then performing 2 x 70% ROOM TEMP  ethanol in order to desalt. This has worked for me many times in the past

2. Mixing your genomic DNA with an equal volume of 70% ethanol and passing down a purification column for a second time (if column purified the first time)

Keep in mind as well if these are new primers, some primers simply do not work even if you follow the rules attached and in these instances simply re designing primers to an adjacent area of sequence can often work

Dear Laurence Hall

Many Thanks

Actually I am using ISSR marker and as a matter of fact I am working on arbitrary/universal primer. what I simply tried to do is just reviewed different publications that are conducted on the same Genera (Thymus sp)   and select those primers for screening  that are highly polymorphic.

After every extraction all the samples were quantified and checked for contaminants using Nano Drop as well as gel Electrophoresis. it is after all this steps that we proceed to the PCR step.

Best Regards

Hello Tesdfaye

Have you ever successfully amplified from these samples with other housekeeping primers to your species ? That way you can separate primer effects from sample variability. I will say nothing further for now

Good luck !!

Dear Laurence Hall

Actually i have amplified  some of the samples successfully.

Thanks,

Dear Tesdfaye

if amplifications were successful from certain samples with certain primers, that implies an issue with primers that didn't work. Sorry I cannot be more specific or helpful than that

Best wishes