is there any reason you can't use temperature alone -the Tm of your oligo minus 5 degrees? That's really the easiest and leaves nothing behind. But yiou can use .1 Molar NaOH and then after 10 minutes or so bring back to the pH you want with1M Tris Cl pH unadjusted. You'll have to figure out by trial how much you need to get the pH to your desired pH

Oops... Use 1M NaOH to adjust your sample to 0.1 M final. This is strong enough to ddenature much lomger pieces of ds DNA as well- such as a full length plasmid.

Actually I am consistantly injecting my oligo sample into a platform at a certain flow rate. There is a sensor in the machine an the system's temperature cannot be set beyond 50, that is the reason why I couldn't melt the DNA at high temperature. I am trying the fix my dsDNA onto the sensor by thiol (-SH) group which has been fixed at 5' ending of one of the ssDNA, so I want to denature it and wash off the other chain. I am worrying if I place the oligo into high base condition, the -SH group connection will be broken.

Is this a synthetic DNA?

Salt, any salt, will stabilize the duplex. Could you please provide more details on the oligo? size in bp, sequence, or calculated Tm?

Not knowing the length of you oligo it is difficult to give a precise answer. Several PCR additives are supposed to reduce the Tm of PCR primers eg glycerol, DMSO, formamide etc. Depending on the length of your oligo the addition of these at 37C-42C may be sufficient to enable denaturation, otherwise the NaOH approach. If you are attaching the oligo to the sensor (silica? fluidic microchip?) via SH groups on one of the strands of the oligos (covalent bond?) then I guess you could try an 5'-3' exonuclease digestion - the attached end should not be available.

Ian

You could try formamide but i don't know good concentrations etc. Its just always in RNA loading buffers to crack up secondary structures... http://nar.oxfordjournals.org/content/24/11/2095.long

But the problem is she wants to denature the ds oligo and then allow cellular RNA to bind it. If you add formamide or even high salt you won't be able to remove it to allow a second annealing.

Thanks to all. I am doing RNA detection, but in this stage, I am using cDNA instead. The oligo is 22bps length. I am able to consisitantly wash the system by PBS, so I would like to try NaOH or urea, if they will not destiry the blocker I used (cysteine).

Thank you, Andrei.

Here's wht some in our group do when attaching DNA by SH group to gold. Since your sensor is under constant flow you can use TCEP along with the ss oligo to prevent secondary structure and electrostatic interactions. These are most likely causing your problem. In addition use a blocking agent along with your oligosomething very small that will bind nonspecifically to the gold, in a ratio that allows enough room between oligo strands to allow flexibility of these molecules and their subsequent availability to your sample virus lysate. Flat surfaces of gold are more difficult to work with than spheres (nanodots) because the perpendicular attachment forces oligos to be too close together, whereas in a sphere the perpendicular angle allows more space as the oligo extends, You may also need to check your oligo for secondary structure.

30 mM NaOH (possibly less, but certainly 30 mM) will denature dsDNA much longer than 22 bp in seconds at RT. Its a great deal more effective than 8M urea or neat formamide (I tested when optimising an ELONA).

Thank you Joan, I talk to one of my biochemistry professor and he also worry about the concerntration of oligo I am using. It can be too high and there is no space to allow target to bind. I am using TCEP to avoid -s=s- formation. Now I am trying to lower the probe oligo concentration. If it still doesn't give a good result, I will try to make it double stranded before the test and use NaOH or Urea to make it denature after blocking.

Thank you Chris. I am trying to use Urea, because in my system there is a cysteine acting as a blocker and I thing NaOH might damage the blocker.

Hao, You need to work on the molar ratio of blocker to oligo. You will need many more molecules of the blocker than the oligo to give enough room for the oligo to renature to your sample DNA and to prevent oligo itself from attaching along its length. I think that will solve your problem without resorting to using ds oligo. I think that will just get you in more trouble. When you add your sample it will need to be in a phosphate buffer to return pH to renaturing conditions.

Joan, I am using 1:10 molar ratio of Cysteine as blocker, but I will increase it to 1:100. I will get back to you if I improve the result with this high blcker concentration.

You mean 1 part oligo to 10 or 100 parts of blocker , Right? Go back over your calculations to measure molar concentrations, to make sure ther's not a simple arithmatic error.

Yes Joan, I meant 1:100 molar ratio.

Helix-coil transition should be possible by pH melting profile of DNA. Pl see the paper appeared in PLoS ONE J. IMJ