I'm doing three different RFLPs with three different restriction enzymes using a single amplicon, each RFLP in different tubes of course as each enzyme test for a different polymorphism.

The enzymes differ in optimal incubation temperature, being 37, 55 and 60 ºC. As is the same amplicon I would like to run a single incubation for the three enzymes. My approach is to set a program on the thermal cycler like this:

37ºC x 4 h
55ºC x 1 h
60ºC x 1 h

My only problem is at which temp to set the thermal cycler lid. I was planning to set it at 60ºC but I don't know how much will this affect reaction on the 37ºC enzyme.

Hope someone can give me some advice.

Thanks

Hello all,

I am new to the forum and posting because I can't seem to find anyone in my lab which does work on qPCR.

I am interested in getting some values of expression level for one target gene. All I want to do is to get a d(ct) value which will represent the level of expression for all my 400 samples so I can see how the expression differs over time and space bz doing some mixed effect model.

I have one target gene an three reference genes, the efficiency of the genes vary by more than 10% so I thought about using the pfaffl method to calculate the d(ct) value. I used the following equation : dct= efficiency target ^ct taget / eff ref ^ dct ref.
However, I realized quickly that there is an issue with my results.
Samples which have the highest expression value (lowest ct value before normalization) ends up with the highest d(ct) values and therefore when I try to plot the values for the manuscript, the lowest expressed samples seem to be the highest.

I hope you follow me, it is a bit hard to explain, all I want to say is that my dct values are kinda reversed and the plot does not make sense and is much harder to explain than it should.

Do you know if I made a mistake or if there is smtg I can do to make my results and graph better?

Thanks a lot everybody

CLaire

Could anyone help me with finding out the difference between using MyTaq DNA Polymerase and Velocity and is there any advantages/disadvantages each has over the other ?

Many Thanks

I have a set of mtDNA Taqman assays for various animal species and I'm trying to validate them. Because this assay is qualitative, I have not been diluting down the DNA after extraction (from muscle tissue) before PCR. The assay has a good sensitivity (~0.05% target in mixed species tissue). However, some of the assays are giving non-specific amplification at around Ct 30-35.

Could this non-specific amplification be due to running too high concentrations of non-specific DNA?

If I dilute my DNA (to 10ng per rxn for example), am I likely to lose my 0.05% LOD?

Thanks.

Since I'm a bit of a novice at molecular biology, I thought I would ask before spending a lot of money on kits and reagents in order to try to sequence whole cDNA using PCR to clone it.

The 2 of the cDNAs I'm after are approx~1.5 kb long. The third one is ~5kb long. I've looked into some high high fidelity pcr kits, and the protocols say that they should be able to amplify >5 kb amplicons no problem.

The problem I have is, how should I design a primer for such a long pcr reaction? I'll start running into primer dimers and hairpins since the reaction is much longer right? The sequences of the whole cDNAs are known (I'm looking for mutations), and I designed my primers to work at the very beginning of the 5' and 3' ends. The problem is that it is virtually impossible to create primers that don't give self complimentary scores <3.0 according to primer blast. Any tips or can I just use such primers anyway? I'm under the impression that primers don't have to be perfect for pcr to work, correct?