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Unsuccessful cloning with one sticky end & one blunt end


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#1 UCI_StemCell

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Posted 25 November 2010 - 03:39 PM

I'm trying to ligate a 1.5 kb insert into the MCS of a 5.3 kb vector and have been unsuccessful so far in isolating a plasmid with the construct of interest. I'm digesting the vector with BamHI-HF (sticky end) & EcoRV-HF (blunt end), and the insert with BamHI-HF (sticky end) and SmaI (blunt end). After digestion I would gel purify and extract both the vector & insert, and use the Quick Ligation Kit from NEB to ligate the two fragments. Following ligation, I would use about half of the ligation reaction to transform DB3.1 bacteria from Invitrogen (I've transformed numerous times w/ this strain without any problems) and the rest of it (~10 ul) I would analyze on a gel & digest with the appropriate enzymes to cut the insert out if ligation was successful. I've tried several different vector:insert ratios and every time I run the ligation product I get bands around ~11-12kb which is not representative of circular DNA, which would run faster, or even linear DNA (5.3kb + 1.5kb = 6.8kb). The bands from the ligation product digestion are also not in the correct size ranges. Finally, post-transformation, I get VERY low transformation efficiencies (1-2 colonies per agar plate) and the isolated plasmid does not contain the insert (which also makes no sense since the ends are not compatible).

I called NEB and their tech support suggested using a phosphatase on the vector to prevent and self-ligation at the blunt end cut w/ EcoRV, however I got the same funky results on the gel and low transformation efficiencies after using Antarctic Phosphatase. My next step is to give up on this and just PCR the insert out of its plasmid and introduce restriction sites that produce sticky ends to make my life easier. Does anyone have any suggestions as to why I'm seeing the results I'm observing from the procedure I have described? Is there anything I can try before I buy primers and a new enzyme for PCR cloning? Thanks and let me know if I need to clarify any of my steps.

I have attached a picture of the gel to help clarify.
G.E. = gel extracted
A.P. = Antarctic Phosphatase

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#2 christy

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Posted 25 November 2010 - 06:26 PM

How did u prepare the insert? Digested from the pGEMT vector?. If the digestion in both the ends are proper and in the vector too, u will get the clone. we did some cloning, we got the clone.

All the best.



I'm trying to ligate a 1.5 kb insert into the MCS of a 5.3 kb vector and have been unsuccessful so far in isolating a plasmid with the construct of interest. I'm digesting the vector with BamHI-HF (sticky end) & EcoRV-HF (blunt end), and the insert with BamHI-HF (sticky end) and SmaI (blunt end). After digestion I would gel purify and extract both the vector & insert, and use the Quick Ligation Kit from NEB to ligate the two fragments. Following ligation, I would use about half of the ligation reaction to transform DB3.1 bacteria from Invitrogen (I've transformed numerous times w/ this strain without any problems) and the rest of it (~10 ul) I would analyze on a gel & digest with the appropriate enzymes to cut the insert out if ligation was successful. I've tried several different vector:insert ratios and every time I run the ligation product I get bands around ~11-12kb which is not representative of circular DNA, which would run faster, or even linear DNA (5.3kb + 1.5kb = 6.8kb). The bands from the ligation product digestion are also not in the correct size ranges. Finally, post-transformation, I get VERY low transformation efficiencies (1-2 colonies per agar plate) and the isolated plasmid does not contain the insert (which also makes no sense since the ends are not compatible).

I called NEB and their tech support suggested using a phosphatase on the vector to prevent and self-ligation at the blunt end cut w/ EcoRV, however I got the same funky results on the gel and low transformation efficiencies after using Antarctic Phosphatase. My next step is to give up on this and just PCR the insert out of its plasmid and introduce restriction sites that produce sticky ends to make my life easier. Does anyone have any suggestions as to why I'm seeing the results I'm observing from the procedure I have described? Is there anything I can try before I buy primers and a new enzyme for PCR cloning? Thanks and let me know if I need to clarify any of my steps.

I have attached a picture of the gel to help clarify.
G.E. = gel extracted
A.P. = Antarctic Phosphatase


Edited by christy, 25 November 2010 - 06:27 PM.


#3 UCI_StemCell

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Posted 25 November 2010 - 07:42 PM

Well it's a plasmid from Origene that contains an NGF-GFP gene that I'm digesting and trying to ligate into the MCS of a different vector. The digestion must have been carried out to completion otherwise I would not have been able to separate the insert from the "mother plasmid" on a gel and gel extract it. I just can't explain the LOW transformation efficiency and the weird gel results when I run the ligation products.

#4 perneseblue

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    Unlimited ligation works!

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Posted 25 November 2010 - 07:52 PM

Open circular DNA does not run faster than its linear equivalent, it runs slower. It is the supercoiled circular form of the DNA plasmid molecule which runs faster. The ligation reaction only produces open circular DNA, with little if any supercoiling.

Given that you can see high molecular weight bands, at least we know the ligase is working. I am assuming that the ligation mix was tested just prior to transformation (no further manipulation were done on the ligation mixture) and thus not a case of losing DNA after desalting (assuming electroporation was the transformation method used)

This leaves two places to look for trouble. The restriction digest and the transformation.

First can you tell us the conditions that you used to digest the vector and insert. In particular the SmaI digest. Could you tell us how the insert was made. Was it by PCR? Was the insert gel purified. How far apart are the restriction sites BamHI and EcoRV in the vector?

Can you tell us about the how the transformation was done. Is this electroporation, chemical transformation? Did you desalt the DNA before hand. Are there any trouble with the cells?
May your PCR products be long, your protocols short and your boss on holiday

#5 UCI_StemCell

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Posted 25 November 2010 - 08:15 PM

Open circular DNA does not run faster than its linear equivalent, it runs slower. It is the supercoiled circular form of the DNA plasmid molecule which runs faster. The ligation reaction only produces open circular DNA, with little if any supercoiling.

Given that you can see high molecular weight bands, at least we know the ligase is working. I am assuming that the ligation mix was tested just prior to transformation (no further manipulation were done on the ligation mixture) and thus not a case of losing DNA after desalting (assuming electroporation was the transformation method used)

This leaves two places to look for trouble. The restriction digest and the transformation.

First can you tell us the conditions that you used to digest the vector and insert. In particular the SmaI digest. Could you tell us how the insert was made. Was it by PCR? Was the insert gel purified. How far apart are the restriction sites BamHI and EcoRV in the vector?

Can you tell us about the how the transformation was done. Is this electroporation, chemical transformation? Did you desalt the DNA before hand. Are there any trouble with the cells?


You are correct in your assumption, the picture I attached features the digested and undigested ligation product prior to transformation.

Both digestions of the vector and plasmid containing the insert were performed in a 40 ul reaction volume. For the vector digestion, I added ~1ug (~11 ul) of DNA to 4 ul of NEBuffer4 and 20-25U of EcoRV-HF and filled the rest of the volume with dH2O. I digested for 30mins at 37C at which point I added 25U of BamHI-HF to the reaction to digest at 37C for another 30mins. After the EcoRV blunt cut, there are only 3 base pairs between the end of the DNA fragment and the recognition site for BamHI, however this should not be an issue as BamHI has 97% cleavage efficiency with only 1 bp from the end. When I performed the phosphatase step, I added 10ul of phosphatase buffer and 1ul of antarctic phosphatase directly to the digestion mix and incubated at 37C for 15 minutes for 5' sticky ends and blunt ends as directed by the protocol.

The digestion of the Origene plasmid was performed in a similar fashion, starting with the SmaI digestion at 25C for 30mins and adding BamHI afterwards for another 30mins digestion at 37C.

Following the digestion times, I split up the reaction volumes into 4 wells per reaction on a 0.9% agarose gel and gel extracted after a sufficient amount of time for running the gel.

After gel extraction, I tried a 8:2 and 8:1 insert:vector ratio (in terms of volume, as far as molecular weight it should be 50ng of vector + ~43ng of insert) in 10ul of ligation buffer and 1ul of ligase from the Quick Ligation Kit from NEB. The DNA was not desalted as I am not using electroporation. I have used DB3.1 non-competent cells from Invitrogen for transformations before and have not had any issues. I make the cells competent with CaCl2 and carry out a normal transformation protocol before plating on LB/Amp agar plates. After overnight incubation at 37C, I see anywhere from 0-2 colonies per plate...

One thing I failed to mention is that I have done this exact ligation in the past with similar results - low transformation efficiency of only two colonies on the LB/Amp agar plate - and LUCKILY both of these colonies contained the insert ligated successfully into the vector backbone. I don't know why the efficiency is so low and why I'm not having the luck this time of obtaining the correct construct!

Thanks for any suggestions you may be able to offer.

#6 perneseblue

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    Unlimited ligation works!

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Posted 25 November 2010 - 09:22 PM

the first thing i noticed is the rather short digest time. It is a lot shorter than I would do. Still uncut DNA should lead to more cells and would not explain why so few colonies were recovered. 100 - 300 per plate would the number i expect.

My suggestion would be to look at the transformation efficiency. I am wondering if there is something wrong with the cells.

Personally I would also cut more DNA for a longer period. Running on such small quantities of DNA gives me the jitters.
May your PCR products be long, your protocols short and your boss on holiday

#7 phage434

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Posted 25 November 2010 - 10:25 PM

CaCl2 prepared competent cells have low efficiency. Also, why are you using DB3.1 cells? Is there a ccdB insert in your assembly? Otherwise, I would not be using DB3.1, but rather Top10 or equivalent. And I'd be buying them, if I were having trouble. If you need to prepare DB3.1 cells, I've done good ones with the protocol on Openwetware here:
http://openwetware.o...competent_cells

#8 UCI_StemCell

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Posted 25 November 2010 - 11:29 PM

CaCl2 prepared competent cells have low efficiency. Also, why are you using DB3.1 cells? Is there a ccdB insert in your assembly? Otherwise, I would not be using DB3.1, but rather Top10 or equivalent. And I'd be buying them, if I were having trouble. If you need to prepare DB3.1 cells, I've done good ones with the protocol on Openwetware here:
http://openwetware.o...competent_cells


There's no particular reason why I'm using DB3.1 cells, they just happened to be available in my lab. I think there are also XL1 cells. I really haven't had any problems with the DB3.1 cells until now. Would you guys suggest trying the XL1 or even another strain of bacterial cells? I think I have two other competent strains from Strategene, but every time I try transforming with Stratagene's competent cells (XL1-Blue & SCS110) I don't get any colonies! I follow their protocol to the exact letter yet I haven't been successful with their bacteria yet..

#9 ElHo

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Posted 26 November 2010 - 09:02 AM

As already told I would suggest to cut more DNA, especially your future insert, and maximize your amount of insert in ligation. Also try ligation over night.




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