Plasmid-Stability? Fragment-Deletion? I'm in dire need to identify and solve - (Oct/19/2010 )

First of all thanks for giving this a look and I'm incredible grateful for anyone trying to help me, since my future may depend on solving this problem.

In short my problem is that I'm trying to clone several 10+ kB constructs, but almost everytime after a transformation I only get plasmids that are too short and appear to be all the same.
It is most definately not a contimination since I repeated it many times and worked as throughly as possible. Experiments I done suggest that the problem is plasmid stability and that somehow specific parts of my plasmids get deleted.

So lets get into this in detail.
I have faced and investigated this problems while working with
1.) Normal cloning of a 3 kB and also a 5 kB Insert in the vector pUWL201 (~7 kB).
2.) Site-specific mutagenesis with phosphorylated primers via blunt end ligation. The construct to be mutated is slightly bigger than 10 kB.

Both constructs mentioned as 1.) were successfully cloned two weeks ago (after around 4 month of work) but I still want to include them since more of the like will follow in the future and I fear I will face similar problems.

Lets first talk about the experiments I've done for identifying the problems with 1.) and which method finally worked. This protocol was performed with both the 3 kB and 5 kB Insert accordingly, so I will just refer to an "insert" in general.:
- PCR worked fine and subsequently was purified via Gel-Electrophoresis and a QIAgen Gel-Extraction kit.
The excision was performed under UV-light. I know this can cause problems but the length of the exposition was very brief and since we don't have an alternative in our lab everyone is using it and it normally never makes any problems.
- Cloning in TopoXL was succesfully performed with the insert. This was done to get easy access to bigger amounts of DNA and also to make sure the enzymatic digestion works properly
- The Insert and also the pUWL201 vector were digested with HindIII and XhoI (buffer2+BSA). For this not the original vector was used but some vector already containing another insert (but still both the HindIII and XhoI cleavage site were present). This was done so one could see both the cut vector and the removed insert on a gel and to properly remove uncut vector (if present). Accordingly the digest of the Insert-TopoXL-construct was seperated on a gel large enough to clearly differentiate between the two sets of bands.
- The Insert was dephosphorylated with Antarctic Phosphatase for 1h at 37°C, following 25 min at 65°C to inactivate the Antarctic Phosphatase.
- As control only vector DNA was incubated with T4 DNA Ligase (and of course T4 buffer) overnight at 16°C and transformed afterwards. When the plates showed no or only 1-2 colonies the sample was used for the next step.
- In a total of 10 uL (containing 1.0 uL T4 buffer, 0.5 uL T4 DNA Ligase) 25 ng of cut vector DNA were ligated with the cut, dephosphorylated insert in a 2:1 ratio (insert to vector) at 16°C over night.
- Subsequently the ligation mixture was dialysed for 1 hour at room temperature.
- 5 uL of the dialysed sample were then transformed through electroporation using 40 uL E. coli TOP10 cells, minimizing physical stress for cells and DNA (e.g. no pipetting up and down to mix cells and DNA) and incubating the cells afterwards with 360 uL LB-Medium at 37°C for 1-1.5 h before putting 30 uL and 300 uL of the sample on different Amp100 LB-plates.
- The transformation yielded, after overnight incubation at 37°C, 10 colonies with the 5 kB Insert (one of which was positive and no cases of uncut vector) and 15 colonies with the 3 kB Insert (one of which was positive and another one which was uncut vector). All other colonies showed shorter plasmids as seen all the different tries before. Both constructs were subsequently sequenced and seem perfectly fine.

Ligation was tested through making 20 uL ligation sample accordingly to what was mentioned above. This was done once with insert+vector and once with insert only. This showed that in both cases the bands of the insert (as control the same amount of non-ligated, cut insert was put on the gel also) faded and a (a little bit smeared) new band was formed at the height of a higher kB number (1 kB marker was used as reference).
Based on this I concluded that my inserts formed dimers as side reactions, which was the reason for dephosphorylating them. Thus I hoped to achieve a higher efficiency in the ligation process.
An accordingly performed experiment with dephosphorylated, cut insert showed that the sample with insert only still featured an intense band of the original insert and almost no dimers. The sample with both insert and vector looked similar but formed a vague smear at a higher position (for this a 30 uL ligation mixture was prepared from which 20 uL were put on the gel, while the remaining 10 uL were used for the transformation which finally yielded both my constructs).

One has too remember that I tried this for around 4 months (while working on other things on the side of course) and each and every other try ended most of the times in plates with (far) more colonies than mentioned above, but yielded only plasmids at around 3 kB on the gel, which weren't cut with KpnI (which was the digestion enzyme I used to varify the identy of my targed construct). This led me at one point to suspect that something happens after the ligation and prior to the plasmid preparation.
When I retransform my construct (or any other construct for this matter) I get a huge number of colonies (which shows my cells are definately competent enough) and never got a single instant where I didn't get the plasmid I was looking for. Lets keep this in mind, I'll come back to this later on again.
Prior to the successful cloning I tried ratios from >6:1 to 1:1 and several different ligation conditions (NEB FAQ suggests lower ratios when trying to ligate bigger inserts and doing the ligation overnight at 16°C)

Now let's talk about what I've done concerning 2.)
- PCR was performed with Phusion-Polymerase, the diluted 10kB construct as template, adding the dephosphorylated primers (and anything else needed) and worked just fine. Purification was performed as mentioned (yielding a good amounts of DNA). After eluating with EB-buffer a 1 hour digest with DpnI was performed to remove remaining methylated template. The digest was performed at 37 °C in buffer 4 and was followed by incubating 25 min at 65°C to inactivate the DpnI.
- Ligation was performed with 8.5 uL DNA, 1.0 uL T4 DNA Buffer and 0.5 uL T4 DNA Ligase at 16°C overnight. (2h at 20°C was also tested at one point)
- Transformation was either performed directly afterwards (using 1 uL undialysed sample) or after dialysis with 5 uL sample (both lead to plates with a sufficiently number of colonies). Again electroporation was used (see above) and after 1-1.5 h of incubation at 37°C, 30 uL and 300uL were put on Chloramphenicol-17 LB-plates (17 instead of 34 because it is a low copy plasmid. The overnight cultures for plasmid preparation contain Cm34 and always grew nicely).

I did this often and prepared, until now, probably at least around 150 if not 200 plasmids. In probably 99% of the cases I get the same plasmid around 4 kB (once in a while I get plasmids which are heavier but still are too small).
The test digestion is performed with HindIII (Buffer2) which should lead to a band at 1273 b and ~9000 b. And here is the kicker which makes me pretty sure that my main problem is the plasmid stability directly after electroporation:
I ALWAYS find the 1273 band. I compared it with a digest of the template and the supposedly 1273 b bands come at EXACTLY the same spot. I tried to sequence it but apparantly the region where my primer should bind is missing.
I did an additional experiment where I took one ~4 kB plasmid, one ~5.5 kb plasmid and the template plasmid and digested it once with EcoRI and once with EcoRV. EcoRI cuts at 5, EcoRV at 4 sites which are predominantely located on the gencluster of this plasmid.
The 4 kB plasmid in both cases only gets linearized. The 5.5 kb plasmid gets linearized by EcoRV and is cut into two bands by EcoRI. One band at around 2250 b is exactly on the same spot as a band from the template-digest while the other one is slightly above 3.0 kB and slightly under a ~3.5 kB band of the template-digest.
In theory I can build a construct which can satisfy my data rather good in two ways.
a.) Just removing the gencluster of interest
b.) cutting it with EcoRI and religating the fragments with 3.5 kB and 2.2 kB. Thus I also eliminate the gencluster, while keeping the resistence

gen. This construct just has a few houndred bases too much in the 3.5 kB fragment (should be around 3.1-3.2 kB).
I took a look in some lab journals of my colleagues and could also see similar findings, but never to this extend (normally most of the times a few of the plasmids were the correct ones, which lead them too neglect any similarities between the correct ones and the wrong ones which also featured some similar bands). This this problem apparantly is nothing new... but if a cloning works nobody is caring about what caused the wrong plasmids prepared alongside the right ones. Only poor souls as myself which face the same disappointing gels again and again, start to look into what causes this problems in the first place. As mentioned I don't have that much of a problem with the efficiency of my transformations as rather some side reactions I can't put my head around. This leads me preparing huge amounts of plasmids like an imbecile, but almost always without any results.

Because of this I tried several changes in the transformation protocol, i.e.:
- Increasing the length of the incubation without antibiotics from 1 h to 1.5 h. This was done to give the cells more time to incorporate and multiply the target construct without any outward stress.
- Using XL1 Blue instead of TOP10 Cells.
All of which lead to the same result: Too small plasmids which all feature a 1273 b band when digested with HindIII.
Still I have no problems with retransforming constructs which are around this size.

I'm starting to get desperate since I have no more ideas of what I can change. I even start thinking that the cloning of 1.) succeeded out of mere coincidence and am afraid that this is the only way to go. Just relentlessy repeating the same experiment over and over again, until fortune shines upon me on one single day...

If any more information is needed about how an experiment is performed, I'll gladly add it. If anything written is incomprehensible, I'll gladly look for other words to describe it. If anyone here is able to help me solve this problem, I'll probably just be very glad myself.

maybe can you give some information on the sequence of the insert you are trying to clone? ...does it contain repetitive sequence elements like direct or inverted repeats? From what origin it is (human, bacterial, viral)? What E. coli cells do you use for cloning?

Maybe you can provide that information and maybe then someone can help you!

Best regards,
p

P.S.: Do not get desperate ...this is molecular biology ...better get used to it

pDNA on Tue Oct 19 20:51:23 2010 said:

The inserts are in general from bacteria. The aforementioned ones from the ordinary cloning are from Streptomyces HK1 and Streptomyces sviceus. The one I used in the mutagenesis experiments is a modified E. coli plasmid. Generally I only work with prokaryote DNA

Is there a way to easily check sequences for repeats besides checking in manually? I could also send you the sequences as CloneManager or textfiles if you this would help.

For cloning I use mainly E. coli TOP10, but also once tried E. coli XL1 Blue with the same results.


P.S.: I am used to it, but my professor thinks differently of molecular biology which could prove to be a problem in the long haul

you can use this two online tools to find either direct or inverted repeats

You can also feel free to read this ...since it references some very good papers and gives you a good overview what could be the reason for your instabilities.

Hope this will help! If you are stuck again let me know!

Regards,
p

By the way:
Anyone else here who had similar problem?
In my lab only one other person had this at a point and also solved it by pure coincidence, so he couldn't give me any advice to solve my problem at all.

Molecular cloning can be one of the most rewarding and one of the most frustrating things you do in the lab. I had problems cloning too (be it mammalian genes) and thanks to the folks on this site and a use-whatever-you-need-just-get-it-fixed from my supervisor, it's now on track again.

I hope we are able to get your problem solved.

- Did you check your competent cells to see if they don't contain a plasmid that could yield a 1273bp band? Same for epps/cuvets and other things used in your experiments (just walk some competent cells through the process like you normally do except don't add DNA).

-EDIT: I forgot to mention this. If you have UV trays with high intensity UV, and you have a band with a decent DNA concentration, you can put the UV shield (that you normally place on your head or between you and the tray) on the UV tray, and visualize your gel on top of it. The UV shield will filter out 95% of the UV, but a small amount passes, just enough to see a band -if you turn of the lights!-. This is very efficient in protecting the DNA from UV, but works only if you have enough DNA. When your done cutting, visualize the gel afterwards in the normal fashion to see if your bands are not contaminated with too much uncut material (which sometimes happens).

- Is there anyway your could try and clone your insert in steps? Say it looks like
EcoRI<-----------------------------------------------------> KpnI 10kb
EcoRI<-2kb->HindIII<-2kb->NotI<-2kb->BamHI<-2kb->ApaI<-2kb-> KpnI

Then you need an MCS in your backbone like this
EcoRI-HindIII-NotI-BamHI-ApaI-KpnI (make sure these are unique sites!!)
You then clone first your Eco-Hind fragment in the backbone, do mini, maxi
Then clone the HindIII-NotI fragment, mini maxi
Then insert the NotI-BamHI fragment, mini and maxi

Skip the maxi if you have pure mini's.

You get the idea. Instead of once inserting a 10kb fragment you insert 2kb fragments with sticky ligation. You only need to make a custom MCS for your backbone.

There are also some other, general measures you could take if weird stuff happens to your construct. (in my case recombinations with Adenoviral vector):
-Grow your plates /minis at lower temperatures (you could try 30C, RT (even better).)
-Grow in rec- strains
-Grow in different host <-never tried this

hematopoietry on Wed Oct 20 09:50:03 2010 said:

Yes I checked once after preparing the batch. Since I use them generally for all my cloning experiments I'm pretty sure it is no contamination, since e.g. control-transformations of cut vector without DNA normally tend to have no or only 1-2 colonies on it. In contrast most of the times doing a transformation of a regular ligation yield lots of colonies, but never the ones I'm screening for.

I could try this, but then again I really don't think the UV is what causes the problems since it is the standard procedure here in our lab and I`d guess it would rather lead to mutations than deletions. But I'll keep it in mind, thanks.

Sadly I need to clone the whole cluster (either 3 kB and 5 kB). Once it was tried to put all the single genes seperately in a pCDFDuet vector but this resulted in a much lower expression rate compared to cloning the whole Cluster.

- Yes, I read at another place about the possibility to incubate at lower temperatures to increase the plasmid stability. This will probably the next thing I'm going to try out.
- Where can I check if my strains are rec- and what are commonly available rec- strains I probably could my hands on?
- Growing in a different host would be an idea, but one I can't realise since we almost exclusively work with E. coli. The one major exception is that we just started working with Streptomyces expression strains. But these grow very slow and we have to clone our constructs in E. coli to have the very least chance to get transformants (as a matter of fact both constructs described in 1.) were made for this purpose).

You can determine recA +/- status on most strains here:
http://openwetware.org/wiki/E._coli_genotypes

Your strains are already recA-.
For stability, I would recommend several things:
* Clone into a more stable strain, such as SURE or STBL strains (See the genotype page)
* Grow at lower temperatures (30C or lower)
* Switch to a lower copy number plasmid (pBR322 or pSC101 or P15 origin, e.g.)

If you can post the (fasta) sequence file, it would be useful.

TGS on Wed Oct 20 11:08:15 2010 said:

Ah, I knew the explanation was a bit to brief. I'm sorry for that. I didn't suggest a permanent alteration of your 10kb insert sequence, but a sequential cloning method that yields your desired vector with the exact same insert as you are trying to clone now. I drew you a picture detailing this cloning strategy(see attached picture "cloning strategy.png").
The first thing you do is ligate a custom MCS (=multiple cloning site)into the vector you want to clone your fragment in. You make this MCS by annealing oligos, then ligating annealed oligos in the vector you want to have your insert in. Notice the order of Restriction sites in the MCS, they match the order of restriction sites in your fragment and the two outer most restriction sites are the one your fragment will be cloned in.
Then, step by step, you ligate each fragment of your insert into this vector, by first obtaining all the fragments through the appropriate digestions, then ligating them one by one into your vector as shown in the picture. Because these are all sticky ligations, with relatively small fragments, they are easy and will not alter the sequence of your insert.

This is a pretty elaborate strategy and should only be used as a last resort. It can also be used to debug the proces, showing you at which step strange things happen.

phage434 on Wed Oct 20 13:10:47 2010 said:

- I fear we don't have SURE or STABL strains availbale and I don't think they will be bought because of me. I'll just have to continue to work with TOP10.
- I'm just dialysing a ligation of the mutagenesis experiment and will try with incubation at 30°C (both after electroporation and when on the agar plate)
- That's also not an option because we need this specific vector as shuttle betwenn E. coli and Streptomyces
- I added the sequences in a .txt as attachment (directly posting would make this thread unreadable):
pSHK1 pUWL201 and pSsviceus pUWL201 are the constructs of 1.), pTUC is the construct which I'm trying to mutate and pUWL201 is the orginal vector

hematopoietry on Wed Oct 20 13:16:07 2010 said:

Ah ok, I understand. Yes, this would be a possibility but I would really be the last of resorts. I'll keep it in mind!
BTW my inserts are only 3 or 5 kB. 10 kB is the size of the plasmid where I want to make a Site-directed mutagenesis, so there don't have this possibility anyway.

Edit:
Looks like the attachment didn't work. Anyone who wants to take a look at the sequences can just write me a private message with his email address and I'll send the sequences via email.