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My battle with BamHI


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35 replies to this topic

#31 phage434

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Posted 18 July 2014 - 06:00 AM

I've just re-read these postings. It seems as if the original poster and several of the followups assume that restriction digestion should go to completion, and that there should be zero background from transforming cut plasmid. In my experience this is never the case. There is always significant residual background, which can be reduced, but not eliminated. If you need to reduce background, then I would strongly recommend using a different method. Some suggestions:

 

* PCR amplify your vector, purify, and cut with enzymes of your choice plus DpnI to get rid of (some) of the template. Add as little template as possible.

 

* Use a visible marker for selecting correct clones (LacZa e.g. with blue/white selection)

 

* Switch antibiotics for your correct construct (Assembly of BioBrick standard biological parts using three antibiotic assembly...by R Shetty - ‎2011 -- Methods Enzymol. 2011;498:311-26)



#32 Trof

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Posted 24 July 2014 - 06:35 AM

We use BamHI from Invitrogen, routinely digest for just an hour in 30 - 50 ul volumes at 37C, and always gel purify our digested vector and insert segments through an agarose gel with guanidine HCL added as UV protectant before ligating, but these don't seem to be related to the problems you're having...


I just got interested in this part. Do you have any protocol for the guanidine HCL loading and it's function as UV protective?
Because by chance I'm doing a restriction digest for cloning tommorow (and yes, with BamHI as a matter of fact) and since I do not clone much I'm looking for any tips how to increase efficiency.
My colleague had serious problems with cloning after gel extractions, until he started using a different UV, that has separate UV-A,B,C UV tubes. When he started using UV-A or B only for cutting out of gel, the cloning started to work.

BTW, as I say I didn't do much of a cloning, and I routinely digest in 20 ul reaction volume (all are NEB enzymes). I have never had any problem with BamHI cuts, but I have with other enzymes, and in several cases increasing the reaction volume to 50 ul or decrasing the volume of DNA added, improved the restriction greatly. Some of RE are specificaly sensitive to salt concentration changes, as it seem. We have several incubators set on 37, on longer restrictions I put the tubes in there, as there is no temperature difference between the bottom of the tube and the top, it decreases water precipitation on the tube lid and keeps the volume stable.

Also in cloning, my boss swears on phenol:chlorophorm purification of all fragments, to get rid of nucleases. He thinks they are the worst enemy for the fragile overhangs. But on the other hand, in his days, cloning was set up on a bench and not in the hood (where all the sensitive reagents and ultrapure water is now only oppened).

Our country has a serious deficiency in lighthouses. I assume the main reason is that we have no sea.

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'Normal' is a dryer setting. - Elizabeth Moon


#33 phage434

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Posted 24 July 2014 - 08:02 AM

Blue light transilluminators eliminate the problem of UV DNA strand breaking. I'd strongly recommend that you switch to using them. They work with virtually all of the common dyes (ethidium bromide, syber safe, syber green). Alternatively, eliminate gel purification as a step in your cloning. There are only a few times now that make it necessary.



#34 Trof

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Posted 24 July 2014 - 11:52 AM

Thanks for the tip, but new transluminator is not an option. We have two and unless the one with the camera breaks unrepairably, we won't be buying new one.

 

As for the gel extraction, I think you refer to using the PCR approach for both vector backbone and fragment so that you don't need to separate. I'm still a bit concerned about the possible errors created in the vector, but at the moment I'm using Phusion PCR for the insert anyway, so the whole idea is more familiar.

 

But for that I would need to buy other things and primers, and I don't have much time (and money) actually. I can do that as backup option if the money was not concerned, but I now prefer tweaks that I can fit into my current approach (which is gel purify both vector and insert and thus limit the damage done by UV). 

I did clone from gel extracted before, I got what I needed at the end, but not that there were many colonies in some cases.. if I recall, there were 4 on the whole plate (all positive).

 

I would like to play with it and explore possibilities, but as far, I get to cloning (TOPO cloning I don't count, that just works)  only at times when the deadline is... well quite deadly.


Our country has a serious deficiency in lighthouses. I assume the main reason is that we have no sea.

I never trust anything that can't be doubted.

'Normal' is a dryer setting. - Elizabeth Moon


#35 seanspotatobusiness

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Posted 07 August 2014 - 02:40 AM

Regarding the evaporation of water, for 37 C incubations, I use the agar plate incubator - the whole tube is then at 37 C and I don't get condensation on the tube surface. I'm not sure why this isn't done by most people.



#36 -AD-

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Posted 20 July 2015 - 08:25 AM

It is the M- F-ing supercoiling!!!

 

During my PhD I successfully used BamHI a lot & found it was very forgiving (i.e. lazy massive over digests over night at 37oC with no problems during subsequent cloning).

 

However, having moved & started a Post-doc, my BamHI digests suddenly stopped working. I was performing multiple BamHI digests & still getting 100s of colonies on my negative control plate (simply the purified vector without insert, ligase or even ligation buffer). It was a small comfort to find this forum and that a minority of people had be experiencing similar problems. However, I tried the suggested comments to no avail (i.e. being very careful about over digestion/volumes/etc)

 

When I ran my digests out on a gel, they ran as a single band (as expected). The uncut vector runs as two bands: Predominantly as an upper 'relaxed' band (running at higher MW than cut band), with a small amount of supercoiled plasmid, which unfortunately ran at same MW as my cut plasmid. Following BamHI digestions, the former band completely disappeared suggesting a good digest, however I couldn't be sure the supercoiled plasmid was being digested.

 

So what I did was gel purify my plasmid pre-digestion, ensuring only to extract the bands corresponding to the 'relaxed' form. Then I digested with BamHI, treated with CIAP & purified (NB: I simply column purified here, I didn't gel purify a 2nd time). 

 

100 ng of vector treated this way yielded no colonies (I did it side by side with 100 ng of vector not pre-purified and this yielded 100s of colonies as before).

 

I concluded that the supercolied plasmid was resistant to BamHI. 

 

Now, I know from my PhD that BamHI is not so ineffective against all super coiled plasmids, clearly the vast majority of people have no problems performing BamHI digests on plasmids. I only have a very basic understanding of supercoiling, but I imagine that there must be a minimum number of kb to accommodate a writhe regardless of twist. I propose that if you are unlucky enough to have a plasmid that just isn't large enough to accommodate an additional writhe (& therefore retains the maximum amount of tension on the DNA) BamHI (& possibly other enzymes) have difficulty cutting.

 

I am sure people who are better versed on supercoiling than me will have a lot to say about this. However, all I know is that gel purifying my plasmid pre-digest completely solved this problem.






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