sequantial digest - (Jun/18/2014 )
bob1 on Thu Jun 19 21:19:31 2014 said:
Ok, in that case just PCR clean between digests and do a gel extraction after the second digest. The first digest should give you only one band as it is a single digest.
The first digest: it gave 2 bands! Very close next to eachother. Is this uncut DNA vs Cut DNA? I suppose the higher band is the cut DNA?
@phage434: Indeed, I will hardly see it if the band is removed or not. this brings me to the my other question. How can I know its the right plasmid I will have to transform the cells with?
There is 1 restriction site in the part I remove, but not sure this will help to see if I have the correct plasmid. Or would I simple see it as uncut plasmid vs cut plasmid on a gel? (as I saw something similar now from the first digest).
bioke on Sat Jun 21 08:08:17 2014 said:
bob1 on Thu Jun 19 21:19:31 2014 said:
Ok, in that case just PCR clean between digests and do a gel extraction after the second digest. The first digest should give you only one band as it is a single digest.
The first digest: it gave 2 bands! Very close next to eachother. Is this uncut DNA vs Cut DNA? I suppose the higher band is the cut DNA?
Maybe - did you run a control of uncut DNA on the gel? That should show you if the band ispartially uncut or not. Another option is that there is more than one restriction site in the plasmid.
bob1 on Sat Jun 21 10:38:28 2014 said:
bioke on Sat Jun 21 08:08:17 2014 said:
bob1 on Thu Jun 19 21:19:31 2014 said:
Ok, in that case just PCR clean between digests and do a gel extraction after the second digest. The first digest should give you only one band as it is a single digest.
The first digest: it gave 2 bands! Very close next to eachother. Is this uncut DNA vs Cut DNA? I suppose the higher band is the cut DNA?
Maybe - did you run a control of uncut DNA on the gel? That should show you if the band ispartially uncut or not. Another option is that there is more than one restriction site in the plasmid.
Yes!
And thats the weird thing:
The control DNA was (at least this is how it looked) located "between" the cut sample!
(but the control sample was very thick..., so not too clear, I also could not take a picture, will need to do it next time).
And no: there is normally only 1 of that restriction site (according to the sequence I have at least)
bob1 on Thu Jun 19 10:45:15 2014 said:
No, just an ordinary PCR cleanup will do between digests - save the gel extraction for the final step if it produces two or more largeish (50 bp or more) fragments, as smaller than 50 bp will pass through a PCR cleanup kit column.
You could also just do a simple ethanol precipitation, but you can lose a lot of DNA this way if you don't have much or large fragments.
Just curious, when you do the PCR cleanup in between: do you bother to heat inactivate the enzyme? I think its not needed because you will "lose" the enzyme during the clean up (will go through the column).
Or is this incorrect?
You don't really care here. If the enzyme is active during the second digestion, so what? The only difficulty would be if it showed star activity (unlikely). You do need to disable/eliminate the enzyme prior to ligation, assuming you are recreating the cut site. If not, you can do a simultaneous cut/ligate reaction, cycling betwen 37 and 16 if necessary.
phage434 on Mon Jun 23 17:41:58 2014 said:
You don't really care here. If the enzyme is active during the second digestion, so what? The only difficulty would be if it showed star activity (unlikely). You do need to disable/eliminate the enzyme prior to ligation, assuming you are recreating the cut site. If not, you can do a simultaneous cut/ligate reaction, cycling betwen 37 and 16 if necessary.
Recreating the site? Not sure what you mean.
I cut with 2 restriction enzymes that give overhangs that match. So I can "close" the plasmid after removing the piece I want out.
So I could cut with the second RE and at the same time ligate it with T4 ligase? (assuming T4 ligase also works in the RE buffer)?
Or?
Yes, although I wouldn't recommend this except for routine procedures that were well tested. The enzyme will only be active on cut sites -- and you are (by ligating a non-identical site) destroying that site, even though it is ligated.
I never tested it, so I guess its best to play it safe and do it seperatly.
I also wonder: the buffer for the T4 ligase is not the same as the one for the RE. Now: the buffer of the ligase also contains ATP , the RE buffer does not, so I would need to use the ligase buffer anyway, so not sure how I can use them both? Or just do the cutting in the T4 dna ligase buffer? Or mix both buffers and add les water?
(I am thinking on trying it, but the first time I'll just do it in seperate reactions).
A second question: when doing the second digest, I have to clean up the DNA again, now: I already lost 50% of my DNA (initial concentration/2 now).
Is this normal?
I think that after a second ethanol precipitation and a gel clean up, I'll also lose a lot and I am worried I end up not having enough!
phage434 on Tue Jun 24 14:17:12 2014 said:
Yes, although I wouldn't recommend this except for routine procedures that were well tested. The enzyme will only be active on cut sites -- and you are (by ligating a non-identical site) destroying that site, even though it is ligated.
You should be able to achieve < 50% loss on purification. We get more like 80% of initial product with ampure bead purification. Even a column should do better than 50%.
You're right, the ligase buffer needs ATP. But ligase is active in most buffers (avoid very high salt). You can just add a bit of ATP and go.
Ligation is near optimal around 20 ng of vector in a 10 ul volume, with equimolar amounts of insert. You don't need much DNA to ligate, and quality is much more important than quantity. Too high a DNA concentration leads to ligating inactive multi-mers rather than transforming circular fragments.
Ok.
20ng should be possible to archieve I hope.
The buffer I need is the cutsmart buffer from biolaps.
It contains 50mM potassium acetate
20mM tris acetate
10mM magnesium acetate and 100µl/ml BSA.
I am assuming the salt concentration is not too high?
So I can just add some ATP?
phage434 on Tue Jun 24 16:56:46 2014 said:
You should be able to achieve < 50% loss on purification. We get more like 80% of initial product with ampure bead purification. Even a column should do better than 50%.
You're right, the ligase buffer needs ATP. But ligase is active in most buffers (avoid very high salt). You can just add a bit of ATP and go.
Ligation is near optimal around 20 ng of vector in a 10 ul volume, with equimolar amounts of insert. You don't need much DNA to ligate, and quality is much more important than quantity. Too high a DNA concentration leads to ligating inactive multi-mers rather than transforming circular fragments.