Problems with SgrAI digest, ligation, killer cut and weird bands on gel - (Dec/15/2014 )
Hello BioForum community,
This is my first post here so apologies if it’s not perfect. I have a problem with my ligation and I would like to ask for your suggestions, and I will try to provide you with as much info as possible to pinpoint the problem better. It’s a long story, sorry, but I think it is necessary to understand what’s been going on and perhaps you can help me find a completely new approach to my situation, maybe I’m already “too stuck” in this to see something obvious.
So, I have two vectors of different size (5.9kb and 6.7kb) with an SgrAI restriction site where I want to introduce an insert of 1.1kb (actually I have different inserts, but they’re all of the same size).
I first followed a protocol from a colleague for the digest and ligation: linearize with SgrAI for 30min, then run on a gel to isolate the cut band, dephosphorylate with CiP, purify with PC purification kit, ligate.
I usually do my ligations as follows:
- One control (A) contains only the linearized, dephosphorylated plasmid
- One control ( B ) contains the linearized, dephosphorylated plasmid + ligase
- All other plates ( C ) contain the lin. dephos. Plasmid + insert + ligase
Due to the fact that I have several inserts I need to clone into the plasmid, I usually have 4-6 ( C ) plates.
If the colonies on A and B are comparably low, I pick clones from C and check with an analytical digest with EcoRI and NotI if they contain the insert. The restriction sites for these are close to the insert, so I get a very clear distinction whether it is present or not (e.g. 0.5kb without insert, 1.6kb band with insert).
Now in the last few weeks I encountered numerous problems. First, since I need to do so many ligations (several inserts), I quickly tried to find a workaround for the gel purification step because purifying twice causes me to lose a lot of DNA. I would need to load quite low amounts of DNA per well to see a good separation of the bands, so I would need to use several wells for each vector to end up with enough DNA for my ligations, plus an additional well for ladder and undigested control respectively, and run them about 1h to get a good separation…. It just takes a massive amount of time and extra work. (I tried this several times and had different problems, usually I got very low DNA concentration after the last purification step and my ligations simply didn’t work.)
Since SgrAI is said to cause star activity, I tested how long I can digest my vector before I get undesired fragments, and found that even after 1-2h, on a gel I see only more and more of the linearized vector. I also checked the supplier’s information, who claims the enzyme only shows star activity if you use too high DNA and enzyme concentrations for >16h, so I used that as a guideline to set up my digests and I decided to try to “overdigest” instead of relying on cutting the band from a gel. The control plate A looked really good (little undigested plasmid) but plate B looked bad (lots of relegation, so dephos. Didn’t work too well). I still had about 1.5 to 3x as many clones on my C plates so picked them anyway to check if any contain insert. And I ended up picking about 20 clones (from the 30-50 per plate), and found not a single one that contained an insert.
Now the new problem seemed to be the dephosphorylation. My previous attempt was to digest with SgrAI for 2h, then add Cip (I know for a fact that it works in the buffer I used because it worked previously in a different experiment too) for 1h, then purify and ligate (1:3 ratio). This time, after I let the ligation stand for 3h I did a “killer cut”, by adding some SgrAI to the reaction and incubating at 37°C for 30min. The SgrAI should digest only the religated plasmid, keeping the number of colonies on ( B ) low and increasing my chance to find a clone with insert on C plates. At first sight, this worked fine. I got a very good ratio between the B and C plates (4-8 times more colonies on C). But as I picked some clones and checked with EcoRI & NotI, I suddenly saw really weird bands on my gel that I have no idea where they could have come from.
After the digest, I would usually always see a band at 5-6kb (considering only one of the two plasmids now) and a second band at either 0.5 if no insert was present or 1.6 if an insert was present. (I managed to get 4 of my constructs right by chance over the time, but it cost me weeks… but at least because of this I am sure that about the bands!). And now, all of a sudden, I see a big band at about 3-4kb and then several (!) bands, like “double bands” on my gel, one “pair” each at about 300bp, 500bp and another at 750bp. Not in every clone, but in different “combinations” on different clones. After this first attempt to use the new protocol, only a single clone shows the desired band at 1.6kb that tells me it should have the insert, but it also has those “mystery” bands at 300bp and near 1kb.
I have no idea where these bands come from. I wonder if it could be some kind of contamination but I have no idea where it could come from or what it could be (I only work with one of the two plasmids at this time, so this shouldn’t be an issue; and I don’t work on anything else, any other plasmids or anything, at this time).
I also don’t really get it why my control plate B looks so good, giving me chances of like 4 out of 5 that I pick a clone with insert, and I pick 4 at a time and get nothing! This could be a lot of bad luck from my side, but I wonder if I’m missing something else here?
I wonder if the SgrAI killer cut messed up my plasmids, because this is the only thing I did differently in the last two attempts. But I figured that the digested DNA would be less efficient in transformation so after picking clones from my plates I shouldn't find any effects of that on my gel…?
I’m at my wits end at this point. Before I pick more clones from the promising plates of my last try, I’d prefer to know why I got these weird bands, or what could have gone wrong with the kill cut. I would have expected SgrAI to cut maybe a bit less efficiently if anything, because it’s not in its perfect buffer during the killer cut, but I wonder if I got star activity during that… and if so, if it perhaps cut out a large part of my plasmid (which would explain the 3kb band instead of the expected 6kb). Then again, I have never seen EcoRI and NotI cutting my plasmid anywhere but where they were supposed to, so how could I have gotten the weird fragments on my gel?
As you can see, I am a bit desperate after so many weeks of work… it sounded like a simple enough thing to do but I’ve been stuck doing these cloning steps for weeks now, with no significant progress (I got 4 constructs and would need at least 7 more before Christmas, ideally).
Thanks for reading through all of this, and sorry for taking so much of your time.
Perhaps some of you have an idea that could help me out? Any advice, concerning the digests, ligation or weird bands, is highly appreciated!
Hi
About all the weirdo plasmids with unidentifiable bands - i wouldn't worry too much about what it is that you are actually seeing, as it seems to be quite certain that it is not the construct that you were aiming for.
In any case, I like "killer cuts" in general - however especially for a enzyme with *activity i would be careful, as the buffer conditions in the ligase reaction may not be 100% compatible with your restriction enzyme.
About the cloning procedure, CIP is not heat deactivable, so depending on how you proceed after dephosphorylation, you may have an active phosphatase in your ligation reaction, which then would dephosphorylate your insert and therefore making it impossible for the ligase to do it's job. A heat deactivable alternative for CIP is shrimp alkaline phosphatase (SAP). In the very least it saves you time as you don't have to repurify after dephos.
I didn't quite get whether you have a PCR amplified insert or wether you cut it out of the second plasmid ( which likely is due to reading your text a bit too quickly). In case it is a PCR insert I had good experience with phosphorylating the primers I used prior to PCR (T4 polynucleotide kinase, heat deactivable too).
also for difficult to clone constructs i sometimes had good experiences in extending the ligation to overnight at 4°C or 16°C
Sorry if this wasn't too helpful.
Good luck!
Hi!
Thanks for all the suggestions.
You're probably right about the star activity, I could imagine the enzyme does weird things in the wrong buffer, but if it really cuts the plasmid also at non-canonical sites, I wonder how the bacteria were able to take up the "fragments" or linearized pieces at all during transformation.
After I dephosphorylated with CIP I used a PCR purification kit. I read before that CiP is not ideal, but I think we don't have an alternative here at the lab. if the problems continue, and my "B" control plate keeps looking bad, I will try to suggest ordering something else!
You weren't reading to quickly, in fact I didn't mention where I got my inserts from :) But yes, they're from PCR. We use Pfu-X polymerase and mutagenesis primers that add restriction sites at both ends, so after PCR I cut with the corresponding enzymes and should thus get phosphorylated ends. But phosphorylation in general could definitely be an issue here, considering also that I saw so much religation happening. Perhaps also the CiP that we use is not working very well, might be worth trying a fresh one (or like you suggested, a different phosphatase).
I also kept one of my ligations overnight (I transformed with 1µl after about 3h at RT, then put it in the fridge o/n and transformed again the next morning to compare) and I'm about to do the analytical digest for some picked clones today, so perhaps I'm lucky this time. if not, I'll probably go back to the gel extraction method, however I'll try to dephosphorylate immediately after the digest (since I know that at least in its original buffer, SgrAI doesn't show star activity within the first 2h at least), and do the gel purification after that, hopefully I won't lose so much DNA that way.
Anyway - thanks for the feedback!
Do you add 5' extensions past the restriction sites on your PCR primers? You need 4-6 bp extra for the enzymes to cut.
Also, are you purifying your PCR product before cutting? You must, to avoid the PCR enzymes extending and eliminating the cut site.
In general, cutting with single enzymes and dephosphorylating is a difficult method, because dephosphorylation is tricky, and without it the background is ridiculously high. You should use SAP or antarctic phosphatase rather than CIP (I simply throw CIP out). CIP damages ends, and can't be heat killed, so you are always doing extra purifications. Use the minimal amount possible for as short a time as possible (titrate by measuring no-insert ligation results).
In general, ligations are almost never the problem. The problems are either in transformation (which seems not to be an issue here) or with the quality of the DNA going into the ligations.
I would strongly suggest avoiding these problems in the future with two-enzyme cloning, preferably with enzymes that can be heat killed. You can prepare inserts as you are doing with primers having RE sites. But you can also prepare vector in a similar way with primers to the vector sequence. You don't need to be limited by the accidents of whatever restriction sites are present in your vector MCS.
Thanks for your advice, phage 434!
> Do you add 5' extensions past the restriction sites on your PCR primers? You need 4-6 bp extra for the enzymes to cut.
Yes, I did that.
> Also, are you purifying your PCR product before cutting? You must, to avoid the PCR enzymes extending and eliminating the cut site.
I did that too.
>CIP damages ends (...) Use the minimal amount possible for as short a time as possible
This is actually really interesting advice, I would have thought if my dephosphorylation does not work it is better to use CIP for a longer period of time (I usually do 1h because it has previously worked nicely with other vectors). I will definitely consider this!
Two-enzyme-cloning would also mean that I can chose the enzymes in such a way that the insert can only be ligated in the correct direction, right? Is there another advantage?
So in principle I would use two mutagenesis primers for my insert(s) that give me two different restriction sites (with different 5' overhangs) for each end, then cut and purify the PCR product. Then I would have to make a new vector with two "compatible" restriction sites instead of the SgrAI site, preferably in such a way that the insert would already be ligated in the correct orientation. This "new" vector would then be digested by the two enzymes, dephosphorylated and purified and used for ligation. IF I get inserts they should be in the correct orientation right away. Is that correct?
I will ask my supervisor if we have this SgrAI restriction site for a specific reason and if nothing else works, I will probably try this.
Thanks again!