I want to ensure that there is a 1.7X to 1.9X increase in the template per cycle. Is it fine if I set up an identical reaction in 30 tubes and after each cycle keep on removing one tube from the cycler such that the first tube has 1 cycled product and 30th tube has 30 times cycled product? Then check the product on gel and with nanodrop and then estimate the activity?
That would work, yes: if you're taking a sample for every single cycle you should be able to capture the exponential phase, though be aware that what you'll actually see is likely to be "nothing for 20 cycles, then something that increases from background noise to saturation over 5 or 6 cycles, then just saturation for the remaining cycles": i.e. you'll only be getting useful data from 4-6 samples (though by using 30 samples you can at least ensure you DO get those 4-6 that you need).
Also, I'd suspect that nanodrop is not a viable approach, since you'll have tons of primers and free nucleotides in your PCR samples, all of which will mask your actual 'new DNA' signal for most of the amplification. So just stick to gel electrophoresis.
However, what you're essentially doing is "qPCR on the cheap", which seems like an odd choice unless you're unable to get time on an actual qPCR machine?
John, thanks a lot!
I do understand that I am trying to do a cheap qPCr of sorts but thats because I was told that i cannot assess the taq activity in the real time...
Well, as a direct output measure you're not measuring the taq activity in either method.
You're measuring the efficiency of the reaction, which is a combination of primer design, annealing temp, taq binding affinity, taq processivity, nucleotide availability, buffering conditions and so on.
Ideally a qPCR reaction should be set up such that everything except the template is present in excess (so waaay more primers than you need, waaay more nucleotides than you need, etc) so that the only real limiting factor is the template copy number. Under these conditions you should see an efficiency of 2 (i.e doubling each cycle), but if you don't see an efficiency of 2, that doesn't necessarily mean that your taq activity is at fault. It could be crappy primers, or too much salt, or any number of other things.
To estimate the activity of Taq polymearse, you need to set up conditions where the amount of Taq polymerase is limiting. That means, you do a standard PCR with everything held constant, except with different dilutions of Taq pol. Then the band intensity should decline with the amount of Taq pol added, all the way to zero.
Then, you compare that against an authentic standard (= a commercial Taq pol with known activity). For example, if the 0.5x dilution of your Taq pol gives you a band as intense as the 1x dilution of the commercial Taq pol, then you can conclude that yours is twice as concentrated as the authentic standard. Always use multiple dilutions to average out errors though.
I show an example of it on my blog here, towards the bottom of the page: http://www.geneandcell.com/blogs/molecular-biology-methods/9505231-purification-of-thermostable-dna-polymerases
To Optimize the Taq pol ...you have play with its concentration keeping the rest of the components in the master mix unchanged. Then as you had said, you could run the gel and find out which concentration is the best and optimum. Start from the lowest to the highest possible range and do various gradient PCR. This I believe should help.
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