Hello,
First, let me describe the problem I'm trying to solve. I transformed yeast with an antibiotic marker to truncate a chromosome, with the fragment having a homologous sequence on one and and atelomeric sequence at one end. The idea is that it recombines with the chromosome at its homologous sequence, while the other end will "dangle", the telomeric sequence will elongate by telomerase, and so that other end of the chromosome will be lost (truncated). (There's another copy of the chromosome present, so no essential genes are completely lost.)
I did verify by PCR that the marker inserted at the homologous end. However, I want to verify that the other end did truncate. One approach I'm trying is using the exonuclease BAL-31. The idea is that after digestion, an "internal" gene (like ACT1) will still be PCR-able, but any telomeric regions (such as the marker I transformed with, if the truncation was indeed successful) will not be.
However, when trying to calibrate the system with two known sub-telomeric sequences, I keep getting wildly inconsistent results. Sometimes I will get no bands at all, other times the subtelomeric regions will persist while the ACT1 band disappears, sometimes all bands appear with no decrease in intensity after digestion. I'm trying to isolate the source of variability, but at this point it feels like I'm missing something fundamental. (EDIT: I did try to deactivate BAL-31 before the PCR using either EGTA or heating to 95C, and all samples were kept in -20 or on ice when working with them.)
In the literature examples of BAL 31 usage I could find, they usually incubate the DNA in an agar plug, and then run the result in gel - no PCR is done. And maybe that's for a reason? Or maybe I just didn't find the right references?
In any case, if anyone has experience with BAL 31, or can suggest another exonuclease (preferably one that was successfully used in a similar scenario), or perhaps give hints as to why the PCR results are so finicky in this case, that would be greatly appreciated.
Yes, PCR can be used immediately after BAL 31 digestion. BAL 31 is a nuclease that degrades both single-stranded and double-stranded DNA from the 3' end. It is often used to generate a series of nested deletions in DNA fragments. After performing the BAL 31 digestion, you can proceed with PCR to amplify the remaining DNA fragments.
To use PCR after BAL 31 digestion, follow these steps:
1. Inactivate the BAL 31 enzyme: Heat inactivate the BAL 31 enzyme at 70-80°C for 10 minutes to stop the digestion process.
2. Purify the digested DNA: Purify the DNA fragments using a purification method such as phenol-chloroform extraction or a DNA purification kit to remove the remaining enzymes and reaction components.
3. Design PCR primers: Design specific PCR primers that anneal to the regions flanking the digested region. These primers will allow you to amplify the remaining DNA fragments.
4. PCR setup: Set up your PCR reaction using the purified digested DNA as the template and the designed primers. Use a high-fidelity DNA polymerase to minimize errors during amplification.
5. PCR cycling conditions: Optimize the PCR cycling conditions, including the annealing temperature and extension time, to achieve specific and efficient amplification of your target fragments.
6. PCR analysis: Analyze the PCR products using gel electrophoresis or any other suitable method to verify the amplification and check the size of the fragments.
Keep in mind that the success of the PCR after BAL 31 digestion depends on the efficiency of the digestion, the purity of the DNA template, and the design of the PCR primers. Careful optimization and controls are essential to ensure accurate and reliable results.
You might find our publication useful:
Article A polymerase chain reaction-based approach for evaluation of...
Best luck with the experiments.
V.
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