Annealing of oligos to form dsDNA?

I have tried mixing them in 1x Annealing Buffer (10 mM Tris, pH 7.5–8.0, 50 mM NaCl, 1 mM EDTA) at 10uM final concentration for both the forward and the reverse and then putting them at 97ºC for 10-15 min and cooling down slowly either on a thermomixer or on a thermal cycler with a decrease of 1 degree/minute after the 97ºC step. I let them reach 25ºC before storing them at 4ºC. So far when I run the single strand DNA (either forward or reverse sequence) and the "annealed" mixture in a 1% agarose gel the "annealed" sample is migrating more (lower) and produces a smear (possible DNA degradation?)

Hello,

I'm trying to clone oligos into my lentiviral vector. I've been having problems doing so, so I thought about optimizing every step of the process. Is there a way for me to check if the oligos annealed together before ligating them with the vector?

I did 1% agarose gel to separate 2kbp ssDNA and dsDNA. According to some papers, they described ssDNA should run faster than dsDNA, however, I ran the mixed sample after purifying the mixture with a PCR purification kit, and the low mobility signal line disappeared. Does anyone have some experience with this?

Figure description: #1: purified 2kbp ds DNA; #2: 2kbp ssDNA & ds DNA mixture; #3: purified 2kbp ssDNA & dsDNA mixture.

Hello I need help putting together annealing buffer. I was given the following:

H2O supplemented with 10 mM Tris-hCl pH 8.5 and 50 mM NaCl

We currently have 10mM Tris-HCL, nothing higher. Therefore I am worried that when I put together the buffer, the diluted Tris-HCL will affect annealing.

Would my annealing reaction still be okay and should I adjust the concentration of NaCl?

I'm currently working on my fourth try of transformation. Every time the plates have A LOT of tiny colonies (photo attached). Every once in a while there is a larger colony, but when I screen them, they don't contain the recombinant plasmid. The negative control has colonies also, but the plate doesn't have the tiny colonies covering it like the other plates.

I've redone the digestion and still get the same results. I checked to make sure the plates contain ampicillin by transforming bacteria with a plasmid resistant to kenomycin. No bacteria grew when I did this. I've tried troubleshooting, but there are a lot of things I can change, and I'm not sure where to start. Is the ampicillin concentration in the plates not high enough (I've been using 500 uL for 500 mL of agar), am I using too much of my ligation reaction (I've been using all 10 uL), am I plating too much (I've been plating about 350 uL)? Has anyone had this issue before?

Also, when you screen your colonies, what else do you run on the gel? I vaguely remember doing cloning in another lab and when we screened our colonies we ran the uncut vector and the cut vector, but I've only been running the negative and positive controls.

I want to anneal two oligos (length: 60bp) to clone it into a vector. They possess sticky ends for ligation. 

How much oligo should I mix (cc. 100 uM) for annealing?

Is NFW suitable or do I need a particular buffer for annealing?

Temperature of the reaction?

After this how much dsDNA needed for the ligation usually?

Does somebody has a good protocol?

Thanks!

Hi

I'm doing asymmetric PCR to amplify ssDNA, using F:R=1:10 ratio of primers.

I already optimized the PCR cycle, annealing temperature of my PCR conditions, but when I detected my product with 1% agarose gel, ssDNA band detected higher than the dsDNA band. As i know, ssDNA has to be lower than dsDNA because they migrate faster. Stranger thing is dsDNA band from the asymmetric PCR product showed higher than the normal (F:R=1:1) PCR product band. My target product's size is 185bp.

Does anyone tell me what is wrong?

I'm using 1% agarose gel, 70V 30-40min (same as DNA ver. agarose gel - I confirmed that electrophoresis gel condition for DNA did not affect to ssDNA detection)