sequantial digest (View forum version)



bioke

Posted 18 June 2014 - 02:27 AM

Hallo all,

 

I would like to make a digest with BspEI and XmaI , their buffers are not compatible, so I need to do a sequential digest.

 

Now can I just use XmaI in the specific buffer (50mM NaCl) and after this reaction is done just add some of the BspeI buffer (100mM potassium acetate)?

Or the other way around.

I read on the NEB website somewhere you can do a sequential restriction reaction using the lowest salt concentration first and than the highest one.

 

Or are they not compatible at all?

 

 

 


bob1

Posted 18 June 2014 - 01:07 PM

From their double digest table it seems that BspEI has 0 activity in buffer 4 and XmaI has 0 activity in buffer 3, so I would clean up between reactions.


bioke

Posted 18 June 2014 - 09:23 PM

I see.

 

So you would suggest a gel clean up than?

 

Would I not lose a lot of the DNA? I heard sometimes you lose almost 80% of the plasmid DNA?


bob1

Posted 19 June 2014 - 02:45 AM

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.


bioke

Posted 19 June 2014 - 03:30 AM

Ok

thanks for the answer bob1

I'll try that.

 

I am trying to remove a 200bp band from a 10.000 bp plasmid.


bob1

Posted 19 June 2014 - 01:19 PM

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.


phage434

Posted 19 June 2014 - 04:22 PM

I now almost universally use ampure bead cleanup for PCR and digest reactions, which I find efficient and much easier, especially with many samples and a multichannel pipettor. You probably won't see a 200 bp fragment digested from a 10 Kb plasmid on a gel unless you load a LOT of cut plasmid, since the band intensity depends on mass, not molarity.


Bio-Lad

Posted 20 June 2014 - 04:54 AM

For what it's worth, you can do a simple column purification on your first digest and then digest the purified plasmid with your second enzyme.  We regenerate our miniprep columns and make our own gel-solubilization/DNA binding buffer so the cost is negligible over the long term and the results are always dandy!  I'd be happy to share the recipe and protocol we use.


paulO

Posted 20 June 2014 - 08:47 AM

Regenerating mini-prep columns sounds terrific.  I used to do TELT minipreps ( no columns needed ) and am a little embarrassed at having succumbed to the column craze.

 

Please share the protocols!


hobglobin

Posted 20 June 2014 - 10:19 AM

Here's a thread:

http://www.protocol-online.org/forums/topic/6134-reuse-dna-spin-column/?hl=%2Bcolumn+%2Bregeneration#entry19532

and a company offers column regeneration kits:

http://www.applichem.com/en/products/maxxbond/


bioke

Posted 21 June 2014 - 12:08 AM

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).

 


 


bob1

Posted 21 June 2014 - 02:38 AM

 

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.


bioke

Posted 21 June 2014 - 05:10 AM

 

 

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)

 


bioke

Posted 23 June 2014 - 09:38 AM

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?

 


phage434

Posted 23 June 2014 - 09:41 AM

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.


bioke

Posted 23 June 2014 - 12:15 PM

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?

 


phage434

Posted 24 June 2014 - 06:17 AM

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.


bioke

Posted 24 June 2014 - 08:35 AM

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!

 

 

 

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.

 


phage434

Posted 24 June 2014 - 08:56 AM

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.


bioke

Posted 24 June 2014 - 09:05 AM

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?

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.

 


phage434

Posted 24 June 2014 - 10:26 AM

Yes. Read the notes on T4 DNA Ligase (M0202) at NEB. Ligase is active in any of their buffer, when you supplement the buffer with 1 mM ATP.


bioke

Posted 24 June 2014 - 10:50 AM

Ok thanks a lot.

Than I'll try your tactic!

Thanks a lot.

 

Yes. Read the notes on T4 DNA Ligase (M0202) at NEB. Ligase is active in any of their buffer, when you supplement the buffer with 1 mM ATP.

 


bioke

Posted 24 June 2014 - 10:57 AM

Oh, one more thing: you mentioned something about cycling between 37°C and 16°C?

 

I know that for the T4 ligase it should be done at 16°C (but thats colder than our room temperature and thus it will be hard to archieve this, we dont really have a cooling system for this)

 

So I'll just use room temperature.

Is there any specific way to do this switching? Start with digesting for half an hour, than half an hour of ligating... and so on? or ?

 

Yes. Read the notes on T4 DNA Ligase (M0202) at NEB. Ligase is active in any of their buffer, when you supplement the buffer with 1 mM ATP.

 


phage434

Posted 24 June 2014 - 01:53 PM

We run digestions and ligations on a PCR cycler, which can easily snd quickly switch from 16 to 37 and back.


bioke

Posted 24 June 2014 - 09:35 PM

Ah yes!

Thats a good idea.

 

One more question: if you do the digest and ligase reaction together, dont you worry that the first cut (with the first RE) will be ligated again? It can be religated if the second RE does not remove the part I want to kick out of the plasmid.

 

We run digestions and ligations on a PCR cycler, which can easily snd quickly switch from 16 to 37 and back.

 


 


phage434

Posted 30 June 2014 - 06:15 PM

Yes, that could happen. We use it for simultaneous cutting with all of the enzymes and ligation with DNA which will destroy the restriction site(s).