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Questions related to Gibson Assembly
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is anybody working on gibson assembly commercialized by NEB? NEBuilder software also helps in designing primer etc. My question is, software predicted annealing temperature differs significantly for taq polymerase and fusion polymerase. Why is it?
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@Juergen is right. It is important to set the proper polymerase. In addition, the underlying thermodynamic parameters used for Phusion Tm calculations are different from all the other polymerases. It uses modified Breslauer data, as originally recommended by Invitrogen, whereas all other calculations use data from SantaLucia. This difference was required to maintain compatibility with the original Tm estimates provided by the old Invitrogen (now Thermo Fisher Scientific) Phusion Tm calculator. Hope that clears up the reason for the significantly different Tm values.
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Hi everyone, I am doing gibson assembly and I tried 4 times but its not working. I checked my primers and all but everything looks fine. The kit which I am using is one year old. So could it be that the kit is no more functional or it has lost its efficiency?
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I can assure you my year old Gibson kit works just like the new one.  I actually compared them directly, with the same reaction, and there was no difference in colony number or correct clone number.  Maybe something special happened to your kit (freezer defrost cycles?).  
With Gibson, not every reaction works, even if you do everything correctly.  So the most likely explanation is that your new reaction simply cannot work due to sequence-dependent problems.  That's one reason they included a positive control assembly with the kit.  If that works, and still gives you the large number of colonies it used to on day one, then it's not the kit's fault.  
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Hello,
I did a Gibson Assembly and after the assembly I stored them on ice, but forgot to put them in -20 degrees. This was at the end of the day. The next morning I saw that the ice was melted and therefore they havent been on ice but water for the night and part of the morning. When I saw this, I immediately stored them in -20 degrees. Now my question is: are they still good to use?
Thanks
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Definitely you can give it a go! All the best.
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When ordering Gibson assembly primers, should either or both of your primer ends (5' or 3') be phosphorylated?
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Hello. Just trying to help here. In this app note which is New England Biolab's, it is unnecessary to  use phosphorylated primers for Gibson Assembly. Here is the link
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I used Gibson assemblies to create an AAV-compatible plasmid. Given the high recombination rate associated with constructs containing ITRs, I tried to transform my assembly product in One shot Stbl3 (transformation efficiency > 1x10^8) and consistently got no clones. I then tried high efficiency DH5alphas (transformation efficiency > 1x10^9) and I get a lot of clones.
I now transform my constructs in these DH5 alpha cells, expand minimally at low temperatures and miniprep. I then use the miniprep DNA to transform Stbl3 (which works beautifully) and expand with these stabler cells.
As this protocol works for me, I would like to know if other people have problems transforming Stbl3 with Gibson products. Their transformation efficiency is reportedly high, And I found them to be very efficient when I transform from restriction digestion/ligation products.
If useful, my Gibson assembly is made from 2 fragments (Vector + insert), I typically use ~50ng of vector and ~60ng insert (4x excess), and run the reaction for 1h.
Is it typical to use very efficient cells to transform from a Gibson?
Should I try running the reaction with more DNA?
Thanks! 
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There is an incompatibility between the Gibson assembly buffer and that of the Stbl3 cells such that the Stbl3 cells are not able to take up your plasmid and therefore all die under antibiotic selection (if you talk to NEB they will tell you this). I have experienced this problem myself and use ethanol precipitation to clean up my assembly product before transforming into Stbl3 cells. It's been my experience that the NEB version of Stbl3 cells are not really that stable, so depending on the construct, cleaning up might be worth it. Hope this helps.
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I'm planning to join together two large fragments (a 10 Kbp vector and a 6 Kbp insert), and I'm wondering whether it's best to use Gibson Assembly or yeast-based homologous recombination. Has anyone used Gibson Assembly to join such large fragments? What's the largest you've ever joined?
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Thanks Manu. I know the protocol says the kit has been tested for up to 20 Kbp, but other than using specific competent cells they don't recommend anything else for the actual reaction. I'll try increasing the temperature to avoid secondary structures though, as the fragment I'm using has high GC content.
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I did a lot of cloning using the gibson assembly master mix and every time it worked perfectly. I must have done more that 30 cloning with great results. 
However, now I've been trying for the last month to do a routine cloning and I only get clones where the backbone closed on itself. 
I want to clone two inserts (1 of 450pb and one of 900pb) into pcDNA3.1 digested with XbaI and BamHI. I purified my inserts and my backbone on agarose gel. I do the cloning with 75ng of vector and and 3-fold excess of each insert.
I tried to do the cloning reaction with only the backbone ( no insaerts) and I also obtained a lot of colonies. How is this possible? What should I do. I already tried to make a new prep of digested pcDNA3.1 with no sucess.
Thank you
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Hi there,
Obviously there is a problem with the vector double digest :
Have you checked the efficient linearization of the vector with each enzyme individually?
Do you gel purify the double digested vector?
If yes to both questions, you shouldn't get any clone with the backbone alone...
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I'm hoping to use Gibson along with a CRISPR protocol and I was planning on ordering my guide RNA along with the primers specified for the plasmid in one oligo. Would I run into any problems doing it this way instead of using two individual primers and doing a PCR?
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You can usually do that.  Although the assembly is going to be more reproducible if you include slightly longer stuffer sequences that always remain constant.  See for example the Church lab cloning strategy from a single gBlock  https://www.addgene.org/static/cms/files/hCRISPR_gRNA_Synthesis.pdf 
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I'm trying to assemble four DNA fragments into a plasmid, each 1.5-1.9 kbp long. Each fragment has an 80 bp overlap with the next. I'm using the NEB Gibson master mix and E. coli top 10 for the transformation. I have controlled for the master mix and the competency of the cell, no problem, but I have no colony after transformation with my Gibson-assembled plasmid. I tried first with 2 microliters, and then next with 5. When I tried with 5 I actually got a colony (only one) but it seems that the plasmid didn't assemble correctly.
I've read some of the explanations given to others with Gibson assembly problem and it seems that my overlap region might have been too long and that secondary structure formation might have prevented correct assembly. Can anyone advise me on what to do next? Is it possible to tinker with the protocol to get better results?
I'm also thinking about PCR-amplifying the fragments but position the primer not all the way to the edges of the fragments so that I can actually reduce the length of the overlap region from 80 bp to about 30 bp and simultaneously get rid of the predicted secondary structures. But I'm worried about mutations to the DNA sequence. Can anyone advise me on this?
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No colonies does not mean your assembly failed.  It means your transformation process isn't efficient enough to tell.  Even in a completely failed assembly you should have scores of colonies from vector carryover and false assemblies.  What do you mean by you "controlled" for it?  Do these cells give you 100s of colonies for other ligation reactions? What is your efficiency with supercoiled fresh miniprep?  
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How to make a cDNA (pAD) library for Y2H protein-protein interaction. If possible let me know step by step with possible illustrations with practical aspects. 
What is the possibility of using Gibson assembly for this?
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See the attached files
Can gibson assembly mastermix ligate a PCR amplified backbone to yield high background?
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Hi all I am attempting what should be a simple gibson assembly and am having some struggles. My background control plate and assembly plate aren't very different in terms of number of colonies and when I sequence my constructs it seems that the assembly works as the inserts are present within the construct but the rest of the insert is deleted (maybe the bacteria delete it?). I know my gene (luciferase) isn't toxic as I have transformed constructs that contained it into the stbl3 cells I am using before swimmingly. My fragments are 9.5 kb and 1.6 kb (insert). I have done both 1:2 and 1:10 insert ratio and get high background and can't find correct clones. I am curious about my background because its something that is occuring in the assembly reaction as when I transform the same amount of vector just in water I basically get not colonies. Can the gibson reaction cause the backbone to self ligate somehow even if it was PCR amplified and not restriction digested? Can anyone recommend an insert:vector ratio I should use for this size fragments? Any help is appreciated. Thanks!
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No, I don't think it's self-ligation.  I've done a lot of assemblies in my life, and looked for evidence of self ligation and never seen it.  It's going to be carryover.  (the way to differentiate is by using different template amounts) I also saw the same thing you're seeing with the vector control -- an untreated vector control is essentially worthless and never gives you the same kinds of colonies the assembly does.  I have no idea what the mechanism of it might be, but it's very clearly what happens.  Perhaps most of the plasmids in a regular prep are damaged and never show up, but are somehow getting repaired during PCR / assembly.  Or maybe the assembly reagent contains some sort of other transfection promoting compound.  No idea.  But it's obviously so, in my hands too.  The good news is that's really easy to fix -- just make your template plasmid as dilute as possible while still allowing the PCR to work.  Try a few dilutions.  If that's a standard mini-prep at 100 ng/ml, try 1 ul each of 1/10, 1/50, 1/200, even 1/1000 can work with a good polymerase.  Now, none of that means that this will get your assembly to work.  It will get rid of the background.  That may or may not bring positive clones up.  If positive clones don't show up, it would be a sequence-specific problem, as Kathryn explained, and in that case, you're going to need to look for a different strategy. 
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Hi
I am trying to CRISPR knockout single genes. I used the LC2 vector and did a Gibson assembly with my gene of interest. Now, the protocol that I inherited mentions that I should transform the e.coli using a certain method. However the protocol is for a CRISPR library and not a single sgRNA (I just want to knock out one gene at a time, using one guide). I read some paper which did not mention transformation and directly packaged the lenti assembled with the guide in HEK cells using different packing systems. 
Please let me know if the simple way of doing it will work.
thanks :)
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No, not even close.  Gibson gives you way too few molecules.  Look at the numbers -- the lenti packaging protocol demands micrograms, but Gibson operates with nanograms.  
You'll need to clone in E. coli to test whether your assembly worked at all, isolate a correct clone, get rid of all the false assemblies.  And then grow the good clone up by itself, to make enough DNA for lenti packaging.  
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I have an insert of 1200bp with >70% GC-content. I managed to pick it up from the genome with gibson primers and with special PCR using ReadyMix for high GC.
The problem is that after the assembly I do not get any positive colonies.
Is there any special protocols for gibson with very high GC levels of inserts? or any tips that might be helpful.
Thanks
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You can try the denaturation before Annealing the PCR product for very small time or say fraction of time. I do say to check the primers during PCR by putting any control DNA sample which has  interested genes, or check for hybridization of your primer  with template before proceeding with PCR
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Hi everyone,
I have been having troubles with my gibson assembly reaction. 
I have a vector, size 7792 bp and insert, size 4633 bp. 
I have successfully amplified both, gel purified and tried 2 to 3 times to do reactions with 50 ng of vector, ratios 1:1 and 1:2. (for amplification I am using Q5) Unfortunately I had a huge background (colonies with a lot of vector). To reduce it I have tried to treat the amplified linearised vector with DpnI and than to gel purify, but now I have almost no colonies. I have repeated this also couple of times and got the same result. I always do a control with only vector to make sure that transformation itself is working. Before this I was using gibson to insert smaller fragments in the same vector and it has always worked very efficiently. I  also reordered different set of primers but the result is the same. Does anyone have any kind of advice? I am not sure what to change...
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Hi Aleksandra,
Is your transformation control plasmid of the same size (about 12.5kb)? The transformation efficiency could be the main issue. Are you using heat shock or electroporation for the transformation?
Additionally, you could apply the DpnI treatment to the final plasmids after the Gibson assembly prior to the transformation. This worked sometimes better than treating the PCR product before the Gibson assembly.
Moreover, you should reduce the amount of template in your PCRs as far as possible. Try different template dilutions until your PCR stops working. Using the lowest possible amount of template should strongly reduce your background in the transformations.
Good luck!
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Hi,
I'm wanting to build some new AAV vectors using a widely used backbone such as this from the Deisseroth lab: https://www.addgene.org/26976/
I have two questions. First, I want to remove certain restriction sites. I've tried with digest + blunting, but it keeps failing on me. The easier way would be to do site-directed mutagenesis, but I've heard that it's not wise to do that on AAV plasmids because the ITRs are challenging or prone to errors in PCR. What are you thoughts on this? Is this true? I've looked but haven't found any papers doing a site-directed mutagenesis on an AAV.
Secondly, I'd like to do some PCR based cloning, like a Gibson Assembly, and avoid restriction enzymes if I can. Same thing here...can I PCR through the whole AAV backbone? Again, the issue might be the ITRs, but otherwise, I don't see why not.
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Hello Alexander - I performed such mutations on AAV vectors (AAV-GFP-shRNA) from stratagene without any problems using the appropriate Taq polymerase (High Fidelity/Proof reading Taq) and standard PCR conditions with DMSO supplement. The mutated vectors were used to knockdown gene expression in vivo and they worked perfectly fine. Don't worry give it a try.
Best of luck
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I'm planning to clone a considerably large Drosophila  gene (~40kb) from genomic DNA into a vector with an attB site for phiC31-mediated integration into an attP landing site. I plan to use Gibson assembly of overlapping PCR fragments, which can reportedly be used to make even larger constructs (https://www.sgidna.com/ultra_kit.html). 
My question is what I would need to consider in cloning such a large insert. I've been having a hard time finding guidelines on the types of materials (vectors, cells, etc) and methods are recommended for this kind of thing.
Thanks for your help.
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You can even with longer fragments as long as 310 Kb. This paper could be of great help.
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Would like to set up an assembly with one fragment. 
Fragment concentration is 38.8ng/ul for 4.1 kb
Vector concentration is 13.1ng/ul for 6.6 kb 
is it simply 3.8 ul vector (50ng) and 3x excess insert (2.4 ul) 10ul master mix and 3.8ul water? 
Thanks!
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Yes, that's correct. Fragment is 4.1 kb, the 38.8ng/ul is 0.01458 pmol/ul. The vector is 6.6 kb, so 3.8 ul vector (50ng) is 0.0117 pmol. 3x excess insert is 0.0351pmol, which is 2.4 ul. If you're using the master mix from NEB, which is 2x, using 10 ul of master mix and the rest of DI water to make the total volume of 20 ul.
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I am using gene specific forward and reverse primers to amplify an insert (2.5 kb) previously ligated to puc19 (2.7 kb), however what I get is a fragment of 5 kb which is equal to the size of puc19 +insert. This is not my own but my labmate's construct. My extension time is 2:30 min with Phusion polymerase, should I decrease the extension time to 1:40 min to resolve this? My plan is to use this amplified gene and do gibson assembly with a different digested puc19, hence I also did the PCR with overlapping primers but get the same result. I will very much appreciate any suggestions. Thanks.
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Hi,
are you sure that what you see in the gel is not the pcr product but a residue of the supercoiled plasmid that you add as template?
In any case, I would suggest to use one primer that anneals in the insert and one in the plasmid backbone. You can also use universal primers such as M13.
And more, add a negative control. Amplify pUC19 without insert with the same primers set and compare to your pcr samples. I would also load in the agarose gel an aliquot of the undigested pUC19 and an aliquot of the putative pUC19+insert, also undigested. 
Hope it helps :)
Gibson Assembly Question?
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Is it possible that a single cut vector can be re-ligated during Gibson assembly? Thanks (sorry for editing..)
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Hi Sabine, there is no re-ligation of the vector in Gibson assembly, even when only cut with one single restriction enzyme. Such religation is prevented by the exonuclease enzyme in the assembly mix. If you get many negative clones (vector) after the assembly the vector used was not linearized completely. In order to join all fragments and the vector in the right order in the assembly you always need the appropriate overlaps of sufficient length. Hope this helps! Gertrud
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I am attempting to clone a gene and a mutant of the same gene into two vectors. The first is peGFP-N1 and the other is pIRES2-eGFP. I have designed primers to add several different restriction sites to the ends of my inserts, (5'-NheI, BglII, 3' SmaI/XmaI, HindIII (not for ires), XhoI) in all possible combinations the cloning is failing. I get literally hundreds of transformant colonies but not a single one with insert. My no ligase plate, and no insert plate each have zero colonies every time. I have since switched to trying Gibson assembly using the NEBuilder kit, used their NEBuilder assembly design tool design my primers and use the Q5 HF master mix for all of my amplifications. In every case the fragments produces are specific and of the predicted size. When doing the NEBuilder the insert are being fragmented, as analytical PCR using the same primers will fail to reproduce all of the fragments but not consistently between samples. After attempting this a few times I changed my plan to excising the eGFP and ires-eGFP cassettes from the respective plasmids and tried concatenating them with my inserts by overlap extension PCR but following amplification the products are the wrong size (about 2/3 of what they should be). I have also tried moving everything to a third plasmid using three fragment Gibson assembly (NEBuilder again) and this always produces truncated inserts. I have sent a form for technical assistance to NEB and their response was it is probably something wrong with my oligos, although they are produced by IDT and validated elsewhere so I know this is not the issue. I have ordered new primers to increase the overlapping region between the fragments from around 15 nts to 30 nts but am awaiting primer arrival to attempt this repeat. Any suggestions would be greatly appreciated, even if very obscure and outside the box. I have attached a gel of my fragment amplification for the most recent attempts at Gibson as well as the analytical PCR done indicating the products are truncated somewhere, and a test digest showing the same result with a greater pool of colonies. FOr the digests, the eGFP constructs should have two BamHI sites, and the ires-eGFP constructs should have three BamHI sites, but as can be seen there is only one in the eGFP and two in the ires-eGFP indicating one has been lost.
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Hi All,
I finally was able to successfully clone these constructs.  To do so I made a fusion of my protein of interest with eGFP in the correct vector and then generated a megaprimer from the IRES-eGFP cassette and used the QuikChange Lightning kit from Agilent to insert the stop codon and the IRES between my protein of interest and the eGFP fusion.  If anyone has trouble with this and would have interest in the protocol I used in designing my primers and amplifying the megaprimer sequence I would be happy to share it, just send me a message.
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What kind of secondary structure must be avoided? I'm trying to assemble 5 fragments (432bp, 546bp, 2333bp, 3674bp, 4329bp). How should I plan my experiment and design the overlapping sequences?
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Normally only highly G/C rich regions make secondary structures strong enough to resist melting during assembly at 50C.  So avoid those.  If it's over 60% G/C, you want to see what you can do to get around it.  A/T rich is not so bad, if the total melting temp adds up, but if you make them by PCR, those may get difficult.  If possible, aim for 50% balance. 
It's tempting to believe the advertisements that 5 junctions are no problem.  Sure, you can find 5-junction fragments that work, but in reality there's some probability that any one junction will not work.  And if you have 5 of them that compounds against you exponentially.  If you feel lucky, you can shoot from the hip and try all 5 at the same time.  But you should be prepared to go back and figure things out one by one if needed.  
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I am in the process of designing a targeting vector for TALEN-mediated knock-in into the mouse Rosa26 locus. The insert will likely be relatively big (8-12 kb) thus also increasing total vector size. I would like to exclude as much unneeded additional baggage as possible.
What would be the bare essential elements requirement for cloning and propagation of the vector in bacteria before isolating, linearisation and transfection for the final knockin? If I understand correctly, I won't be needing either f1 or sv40 ORI which are often included in preexisting vectors. 
I am planning to build the vector from scratch using modules (such as selection marker, origin etc.) from different template vectors. The final targeting vector will be sequentially assembled using Gibson assembley (NEBuilder HiFi).
So far I am planning to limit the vector to following elements:
Vector backbone
  • pUC origin
  • kanamycin resistance
  • bacterial promoter for kanamycin resistance
  • sv40 early promoter for mammalian G418 selection off of kanamycin resistance
Insert for mouse knockin
  • Promoter + Gene + polyA signal
  • Rosa26 homology arms
Genes to be knocked in will include fluorescent proteins, thus I am planning to reuse them for identification of successfull knock-in cells. This way I can avoid the unnecessary insertion of a resistance gene into the mouse genome.
Is there anything that I am missing or would the minimal backbone suffice?
Has anybody taken a similar approach or is this way too crazy?
Thanks a lot for your input!
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I once made a minimal vector just for propagating plasmid in bacteria, it is ~2kb, only has:
origin
Amp resistance
bacterial promoter for Amp resistance
customized MCS
For your purpose, you can modify from a pEGFP vector.
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Hello everybody, I have a problem with DNA cloning into vector. When I amplified target gene, and then insert this gene into vector by Gibson Assembly, and transformed in DH5a. I got some right colonies. I isolated DNA plasmid and sent sequencing (I used primer in vector for sequencing), I already got right result for this gene.  After that, I transformed in BL21 to check protein expression and enzyme activity. However, this result was not good. I had to do many times and I got the same result. Hence, I had to isolated DNA plasmid in BL21 strain and sent sequencing again. Surprisingly, I got result that my target gene was be converted into Ecoli's gene. So I already made again and some time I got the same result. I don't know, what happened!!!! And I can't explain for this case. Please, could you help me????
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BL21 is recA+, but cloning strains are generally recA deficient. So you might have had recombination between plasmid and chromosome. However, this is unusually rare. Is your cloned gene toxic? Unless you are expressing from  T7 promoter, you might try expressing your gene in your cloning strain or any other recA- strain. 
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Hi there,
I am being in troubles with my Gibson Assembly results.
Now I am telling you the situation. Just you can see below agarose gel picture, the vector ( 22125 bp) was from a home-made plasmid PPV5´BD-GFP double digestion. And three inserts NIb-GFP_2 (2436 bp) + pPA11m_2 (2295 bp) + CP3'UTRNOS_1 (1614bp), which were produced by PCR with primers overlapping around 25 bp. All the fragments  were purified through column, while the expected band of CP3'UTRNOS_1 was cut from the gel.
At first, I tried Gibson Assembly flowing the NEB HiFi Master Mix kit, all the fragments amount were loaded to 0.05 pmol, but with 5 µL 2*Master Mix. After 2 hours incubation at 50°C, I didn't observe any bands with loading 1 µL ligation on agarose gel, even nothing in the positive control. Then I did the transformation in both my expected plasmid and the positive control. Finally, no colony in the palsmid plate, but good colonies in the positive one.
I thought probably there were too many fragments.So I tried two-step Gibson Assembly, that is firstly to do the same procedure in the three shorter insters, later to assemble the product with the big vector. Until now I am writing this question to you, I checked the first three fragment assembly without any bands on the agarose gel. This bad situation really confuses me. I don't know what is the problem, maybe I used a decreased amount of Master mix, however which worked in the positive control. Or perhaps the size of the fragments is too long.
Can anyone who had experience in Gibson Assembly help me with this failure? Thank you very much!
Yu
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Hi Yu,
how did you solve the problem with the gibson master mix? I am having the same problem... would you be able to help me?
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I'd like to do a Gibson assembly DNA cloning with a single restriction enzyme (BamHI) digested vector. Should I dephosphorylate (ie. with calf-intestinal alkaline phosphatase) the vector first in order to prevent self-ligation?
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I generally do not phosophatase treat while doing Gibson Assembly.  In fact in the past I have found using antarctic phosphatase treated vector was not as efficient for Gibson Assembly and I ended up getting more background because some background exists due to uncut vector.  I now almost never phosphotase treat.  I actually found your thread because I was trying to see if anyone has used CIP treated vector that was purified afterwards for Gibson and if that reduced efficiency.  For my Gibson Assemblies, I almost always directly use PCR product added to cut vector that has the enzymes heat inactivated but not otherwise purified.  It works great for all routine cloning and even for some library cloning.  
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Hi everyone,
I'm trying to perform an assembly using the NEB Gibson assembly kit. My aim is to assemble 2 fragments : one of 5690 bp (my gene of interest genomic DNA with it endogene promoter) and my vector backbone of 9947 bp. I followed the protocol, i put about 100 ng of vector and 300 ng of insert for a total volume of 6.2 µL and I incubated for 1 hour. After my transformation there was no colonie on my plate...Whereas in my positive control (provided with the kit) it was ok. Can you help me with it please, maybe you have some good advices?
Thanks a lot!
Julie
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Your forward overlap has 2 restriction enzyme sites (palindromes). NEB recommends avoiding this, and avoiding placing the overlap in a Multiple Cloning Area in general, for this reason. I also see a hairpin near the end (which has to primer extend in the Gibson fill-in):
TTGCATGCCTGCAGG  This hairpins with the following:
GACGT...            The first end nt here is a GT mismatch (the most tolerated). Following this are 4 nt of perfect match. The end of the strand, here,  must anneal, and primer extend to reach the 5' end of the other DNA fragment exposed by the exonuclease, and then the two ligate. Could this hairpin, or the restriction enzyme sites, interfere with the overlaps annealing efficiently? I don't really know if this can happen at 50C. If Peter Weigele is still monitoring this his comment is welcome, or you might call NEB and ask them.
I suppose NEB's builder tool can control one end of the overlap, by controling the length or the overlap, but it can't control where we tell it to start and stop the vector from. Maybe this is a reason to let the tool put the overlap where it wishes, and not forbid it to put insert on the vector primer by selecting the vector as "backbone", as I usually do so that I may reuse the primers and PCR product for future cloning.
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Hello everyone, 
I need suggestions for designing primers for Gibson assembly for assembling 3 fragments and finally circularising it, so I will need a total of 6 primers. I have designed overlapping primers using software like serial cloner. The Tm for 4 primers out of 6 is around 73oC. and overall GC content is 60%. The overlap is kept between 25-30 bp. for example
1. Rev : CTGGTAGGCTACGTGTGTCCTATAGTGTCACCTAAATCGTATGTG
2. For: GATTTAGGTGACACTATAGGACACACGTAGCCTACCAGTTTC
Please do suggest if any other criteria need to be taken care of. 
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Dear Hiba, 
have you used Gibson's assembly kit ??
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Hi, 
Can someone help me please? I am trying to insert a gene fragment of 1200 bp into a 5 kb vector using NEBuilder Hi-fi DNA assembly master mix. I have used 3x gene insert with 40 bp overlaps with vector on each side. Gene fragment and vector (0.06 pmol) was incubated for 1h (or 15min) at 50 degC for assembly, as mentioned on NEB's (or IDT) website. Reaction volume used was 20 ul. Assembled plasmid (1ul) was then used to transform BL21(DE3) electrocompetent cells (25ul) and I did not see any colonies. No colonies were obtained with NEBuilder's positive control either. In agarose gel did not see any band for positive control. I had run transformation control with different vector (pBad) and got colonies with that. So, it seems like assembly is not taking place. My labmates have got assembled plasmid DNA using same master mix stock and thermocycler. I am unable to tease out the problem. I would really appreciate if you can help me troubleshoot it. 
Thanks
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Gibson assembly can be tricky and often requires a few attempts with different tweaks to concentrations to determine the optimum. As you are not getting any colonies from your positive control then there are 2 most likely options, 1) The mastermix is degraded and no-longer working, since your labmates had success this is unlikely but have they had success since you have failed? It is possible it has degraded since it was last used. 2) The competent cells are defective. If competent cells undergo repeated freeze-thaw cycles they experience a dramatic loss in competency. Are you using the cells provided with the gibson cloning kit? Gibson mastermix can inhibit other types of competent cells due to the components within it. 
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Hello,
I am constructing a plasmid with a 1.7kb insert into a typical 3.7kb backbone. My enzymes are SalI and NotI. PCR to amplify my insert went very well, and I have used this vector backbone in many other experiments. The problem is when I pick colonies off the plate for miniprep, they do not yield the insert upon digestion. The ratio of my insert/vector combo to vector only is roughly 70:10. I've gone through 7 colonies from 2 different ligation transformation experiments using the same insert/vector combo, and no inserts digest out of the final product. The size of the miniprep product seems too small to have contained the insert as well. What are your thoughts?
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may be your vector religate without insert integtration
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I'm observing a strange phenomenon when I try to recombinantly produce protein in E. coli. I've tried working with two separate genes (completely different codon usage) but I still see the same point mutations when I insert the first half of my gene into my expression vector, pET28a. I'm inserting the gene step wise by splicing fragments via overlap extension (SOE) PCR because I am unable to amplify the full gene. This procedure has worked with numerous other genes, but for some reason I have been having extensive issues with these two (again, they both code for the same protein). Here's a step by step for the methodology:
1) Amplify and purify two gene fragments using terminal primers and complimentary primers specific for the interior of the gene.
*20 ul Q5 Hot Start master mix (NEB), 20 ul ultrapure water, 0.5 uM primer concentrations, and 1 ng template, optimized amplification reaction conditions w/ 30 cycles.
*Confirm amplification on a gel. 
*Clean samples using a GeneJET spin column (ThermoFisher).
2) Insert first gene fragment into pET28a using SOE PCR. 
*5 ul Q5 Hot Start master mix, 5E-5 nmol pET28a plasmid, 5E-5 nmol clean gene fragment insert, fill reaction to 10 ul with ultrapure water, optimized splice reaction conditions w/ 15 cycles.
*Add 0.5 uM of each terminal primer to the above reaction mixture, optimized amplification reaction w/ 25 cycles. 
*Confirm insertion on a gel. 
*Clean sample using a GeneJET spin column.
3) Ligate linear construct and clean product.
*20 ul purified linear construct, 1x T4 DNA Ligase Buffer, 1x Cutsmart buffer, 10 units DpnI (0.5 ul), 1500 units T4 DNA ligase (0.5 ul), 5 units Polynucleotide Kinase (0.5 ul). All enzymes and buffers from NEB. Incubated @ 37C for 2 hours and then @ RT for 2 hours. 
*Clean sample using a GeneJET spin column.
4) Transform T7 Express Competent Cells (NEB) with circular construct.
5) Perform colony check, culture promising colonies, isolate plasmids, and sequence using Sanger sequencing and ReadyMade T7 Promoter or T7 Terminator Primers (IDT). 
*Colonies checked for GOI using PCR and gel electrophoresis. Promising colonies cultured overnight @ 37C in 15 mL centrifuge tube (12 mL media), 1 mL used to bank cells and the remainder used to isolate plasmid (see link below for procedure). Sequencing performed by an outside party (I am confident in the sequencing data). 
Here are the issues I'm encountering:
1) All samples have had at least one point mutation. 
2) Samples that have multiple point mutations are of the same type (either G to T throughout or A to C throughout). 
3) Patterns in point mutations have been observed between genes with completely different codon usage (for example, I to M at amino acid position 166).
Here are my thoughts:
If I keep repeating this procedure I will (statistically speaking) eventually find a successful mutant. However, the costs of this work/screening are kind of getting out of hand at this point. I would like to better understand why the insertion of these genes is proving to be so difficult. I feel that these errors are related to PCR amplification somewhere within this process, but if this is the case wouldn't the point mutations be random in nature? Additionally, I'm using a polymerase with an extremely high fidelity, so PCR being the issue really doesn't make sense. Why am I seeing patterns in the type of mutations and their location? Any and all suggestions and comments are welcome, and thank you ahead of time for your help
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Are you sure the mutations aren't actually in your template? It may be worth resequencing the whole gene if you are able.
You could also change your cloning strategy and try something like golden gate cloning. May involve an extra step or two but it is pretty easy and relatively quick. 
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Hello
I'm trying for a long time now to assemble two fragments (one is 640bp and the other is 100bp) with the Gibson cloning kit. Both fragments were designed with the NEB assembly tools, but I don't want to put a vector on my reaction - I just need the final 740bp product.
I tried always the combination of equimolar concentrations and reaction times, ranging from the lowest recommended until the highest, and I'm not really sure why it is not working.
After the Gibson I always do a PCR with the forward primer of the 640bp fragment and the reverse of the 100bp one, resulting always in two bands.
Could you help me or have any thoughts in why it is not working?
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Actually Enikő Besenyei you can do PCR after Gibson. I do it all the time for my recombineering work. However, I only allow my Gibson Assemblies to run for 15-30 minutes rather than an hour. I find that having more than 25 bp overlap between the two fragments yields better results (I usually use 30-40bp overlap). Also, before PCR I dilute the GA reaction 1:4 because there are PCR inhibitors in the master mix.
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Hi
I am new to Gibson cloning and am running into trouble on getting the Gibson assembly to work:
I have a 7.4 kb vector back bone with 3 fragments (674 bp, 877 bp, 1195 bp)
I tried to assemble in equimolar conc. with 3x excess or 6x excess of inserts, but I didn't get any colonies. Incubation was for 1 hr at 50C with the NEB Gibson assembly mix When I put the assembled mix on a gel I still see 4 seperate bands appearing on my gel instead of assembly...
Any advice on how to get this to work? 
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Hello Steffie
Gibson cloning can be quite tricky sometimes and troubleshooting it can be fairly painful. Just wanted to confirm, that your vector backbone was linearized with the restriction enzyme that you used in NEBuilder to create the complementary regions in the insert fragments right? Gibson assembly may not always assemble as we like it, but after 1h seeing 4 distinct bands may mean that the design of your overlapping regions in the PCR fragments was faulty, whereby there was no stitching by the polymerase and ligase.
Another reason could be that the assembly mix just conked off. This can happen if frozen and thawed too often. 
At this point, I would recommend to check your overlapping regions and readjust your molar ratios of inserts to maximize probability of ligation. I normally use a 5:1 or 7:1 ration of Inserts to Vector molar ratio when assembling. Hope this helps
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I hope somebody is able to give me suggestions.
After treating a 50:50 mix of fully methylated (FM) and non-methylated (NM) commercial DNA (zymosearch) I amplified the locus I want to study. Once excluded any PCR bias - i.e. by digesting the PCR product with a restriction enzyme that cut only the FM "allele" and confirming an almost equal amount of the two products -  I proceed with TA cloning (pGEM + T4 DNA ligase Promega). The ligation product has been used for transforming DH-5a cells. When we checked colonies (nearly 20) to confirm that the 50:50 proportion has been mainteined, we observed instead a 70:30 (FM:NM). This situation has been already reported (Warnecke et al. / Methods 27 (2002) 101–107) as possible.
Do any of you experienced a similar situation and how have you fixed it? Changing maybe ligase or cells?
I thank you in advance
I'm cloning bisulfite treated DNA, in particular a 50:50 mix of fully methylated and non-methylated commercial DNA (zymosearch) 
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Hi Gianluca
I hope that I have correctly understood
Summarizing, you did a 50:50 mixture of methylated and unmethylated DNA, then you did a PCR for methylated DNA (I assume you used oligonucleotides for the BSP method), from there you cloned and sequenced, am I okay?
Through these methods it would not be possible to observe post-cloning a 50:50 methylation ratio, because there are three steps in cloning that make the whole process random. First, despite confirming that there are the same amount of methylated and unmethylated DNA amplicons, it is random that the ligation in the vector is also 50:50; second, it is also random that competent cells capture half of vectors with methylated and unmethylated DNA; and thirdly, the selection of the colonies is also random, there is nothing to ensure that you choose 10 colonies that having the vector with methylated DNA and 10 colonies that having the vector with unmethylated DNA.
Thus, one question comes to mind, is not it clear to me why you using the BSP method to measure the global methylation of a gene? If you use a mix of methylated and unmethylated DNA from Zymo as a control,  would not it be better to find for the relationship through High Resolution Melting?
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I am following the CombiGEM CRISPR protocol to ligate together a lentiviral vector already containing 1 gRNA to another gRNA fragment in order to construct a double KO construct. The ligation is sticky end ligation. The vector is 9700 bp and insert is 400 bp. My previous 2 ligation attempts have failed. I ligate a room temperature for 2 hrs using standard T4 ligase. After my first failed attempt I cut new vector and insert for a longer time and although it looked better on the gel it failed. I have also gel and column purified my vector and inserts. Any suggestions? Should I try to ligate overnight in a thermocycler at 16 degrees?
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My questions refers to the last step in my protocol which is attached.
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I have had a gBlock fragment synthesised by IDT with pOPINF flanking sequences at both the 5' and 3' terminus' and am trying to clone it into pOPINF. The plasmid was successfully linearised with KpnI and HindIII (both enzymes have been checked) and the linearised plasmid gel purified. The purified concentration was low (20ng/ul) but I also ran a gel to check and the linear plasmid is still there. I went ahead with the clonase reaction using my gBlock fragment (~350bp) and the control insert supplied with the clonase pack (~2kb). Transformation was carried out into Stellar competent cells using both clonase reactions, plus a transformation with linear plasmid and another with the complete pOPINF plasmid. Colonies are present on pOPINF transformed cells, but nothing for linear, control insert or my gBlock fragment.
The problem seems to be with the clonase reaction. I am using 40ng of each linear vector and insert and am incubating at 50'C for 15mins, all as per the manufacturers instructions. I have repeated this 3 times with the same result each time. Is anyone able to offer any suggestions as to what the issue may be? Or maybe something which I would try?
Any help would be very much appreciated. Thank you.
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We use inFusion rather than clonase (as do OPPF), but that should make little difference. Good DNA concentrations are always helpful, so making some fresh linearized vector would be a good idea as well as making sure you have plenty of insert as Mark suggests. If you're using a UV transilluminator to visualize your DNA fragments during gel purification, you are damaging the DNA and reducing efficiency, if so, try a blue light box with GelGreen/SYBRGreen. If you get it right, these ligation independent cloning methods are much more reliable than the old ways.
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Hi there,
I have been trying to clone 3 fragments (~1kb each) into a plasmid backbone (6.6 kb) using Gibson Assembly Mix from NEB (https://www.neb.com/products/e5510-gibson-assembly-cloning-kit), without success though. The minimum overlap is 30 bp.
So far, all I got was 10-20 colonies per plate but when I miniprep them it turns out there's only empty vector. Of note, I don´t get colonies after overnight incubation, only after extended incubation (48h), which I find really suspicious. Because of that, I've also been incubating plates at lower temperature (30ºC/ RT) because I suspected that my insert would be somehow toxic for the cells, but the problem remained.
Here follows some details:
- plasmid was digested with a restriction enzyme, run on a gel to ensure complete digestion, band was excised and column purified (~60 ng/uL);
- inserts were PCR-generated (using Phusion), run on a gel (single sharp bands with expected size) and column purified (~100 ng/uL);
- different insert to vector ratios have been tried: 1:1, 3:1 and 5:1;
- extended incubation time (4h) has also been attempted;
- reaction was used to transform chemically competent cells (provided with the kit);
Some controls have also been performed:
- Transformation with uncut vector: >100 colonies (so everything is OK w/ cells and plasmid);
- Transformation with cut vector: no colonies
So, at the moment we have no clue about what is going on with this cloning! Hope some of you may have an idea.
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Hello everyone,
I am planning to perform Gibson assembly for splicing 3 fragments and finally want them circularised. Previously I was performing overlap extension PCR but that did not work for me, I guess the reason being the size of fragments(>4.0 kb each).
I have designed primers for Gibson assembly. The primers for the 3rd fragment is GC rich upto 75%. 
How should I go about this cloning procedure? Do you have any suggestion?
It would be great if you can guide me through this.
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I'm afraid this is not likely to work.  Gibson's handles high G/C very poorly, especially in the overlaps.  At the very least break it up into individual assemblies, so you can learn from a failure where the problem is.  You may get lucky, but your luck drops exponentially with the number of fragments.  Getting your sequence at all is more important than rushing it.  Also you should probably get comfortable with the idea that this may not be a solvable problem.  Look into de-novo synthesis to make this. 
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Hello.
I'm trying to amplify a double strand DNA with sticky ends (not compatible with any restriction enzyme). I'm gonna try a PCR with different types of primers/oligos but I'm considering using oligos with spacer so I can get the sticky ends a little bit easier, but actually I don't know if it's going to work. Do you have any advice?
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Use the type IIS (golden gate) of restriction enzymes, such as BsaI, BsmBI or BbsI (see https://www.neb.com/products/r0535-bsai, as example) with this enzymes the recognition site (Rs->) is directional and point to the sequence just next to it, which is cut leaving a sticky end with almost any sequence (either up or downstream, according to how you orient the recognition sequence) giving you freedom in the choice of sequences on both ends. Therefore, you can use the same enzyme site in the primers flanking your amplicon, but in opposite orientation pointing inwards (Rs->..GGTCTCN(XXXXXXNN..NNZZZZZZ)NGAGACC..<-sR). When you digest the fragment, each end is cut to leave a different sticky end on each side (for directional clonning; 5'XXXXX(N)nZZZZZ3'). The sites have to match the sequences in your destination vector, which has to have two BsaI sites at the insertion site but this time pointing owtward  XXXXXXNGAGACC<-sR(NN...NN)Rs->GGTCTCNZZZZZZ. After digestion with BsaI, the insert can be ligated with the vector and no BsaI sites remain. Actually, tanks to this, you can do the ligation and digestion in the same reaction at the same time, as any other  pairing of fragments leads to products with BsaI sites and thus gets redigested.
A graphic scheme of this can be found here:
Best wishes
Rogelio
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trying to amplify the backbone of a plasmid, amplifying two fragments that will be used in three fragments Gibson assembly with a protein gene of interest.
I got one fragment amplified but i am having trouble amplifying the second fragment. One of the primers is a forward primer with his tag located at the 5 primer end.
does anyone know if his repeat effectss the primer binding to the DNA or something?
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Also make sure your primer has long enough complementary starch, at least 18nt
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We are trying to clone a plasmid to have cre dependent expression of the protein of interest. However we are a bit puzzled with the primer designing. We will do  it by gibson assembly. We have a plasmid with the sequence of interest in the normal orientation, and now we should clone it as a reverse complement, I believe. So I think that when designing the primers for PCR amplification, the overlapping sequences with the vector destination, should be swapped and complement?
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Hi Sara,
can you be a bit more specific about what a Double Flex construct is?  I wonder if the reason why there have been no responses yet is because the terminology is confusing those of us that want to try to help.  It certainly sounds like an interesting technical problem.
If I have understood the above from the detail already described, you want to generate a construct where the reading frame is reversed and not being expressed until you introduce Cre ; then it will be turned around into an orientation where the reading frame is active?
Do you have a diagram of this?
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I have amplified two fragments 4.0 kb each with overlapping segment. I am trying to join them using HerculaseII enzyme. The procedure I am following is this: PCR 1 (overlap PCR) for 10 to 15 cycles and PCR 2 (extension PCR) with end primers for 30 cycles.
I am not getting desired size amplification 1.e of 8kb. Though below 3 kb amplification is seen. I have also increased annealing time from 10 minutes to 30 minutes for PCR1 but still, no desired amplification is seen. 
I have amplified two, 4 kb segments separately and have also gel purified them using QIAGEN kit. Now I need to join them and make a 8 kb amplicon. And this part is troublesome. 
I am attaching the gel image. Please have a look. The ladder is of 10kb. I am getting multiple nonspecific bands but not 8kb desired band.  
My final goal is to clone this final 8 kb fragment.
Need suggestions. 
Thank you!
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Getting an 8 kb amplicon by PCR may not be trivial; have you checked with a different control (a reaction that would yield an 8 kb or longer amplicon) that your conditions are OK before proceeding to SOE?
Also, why don't you try Gibson assembly instead?
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I have a fusion protein made up of the 3 antigenic region  linked by GPGPG and GGGS linkers. It is synthesised and cloned in pUC 57 Kan vector from third party vendor. I need to subclone it into pCOLD vector 1 and express. where HIS tag will be incorported via this vector.  When I digest and release the insert from pUC I get positive result. But by cloning into pCold and the double digestion fails to release the same insert (blank result).  enzymes used are NdeI and XhoI. What could be the reason that I am not able to digest even after overnight digestion??
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Hi....I did over lap extension  PCR  based cloning of two separate DNA fragment each size of 0.8kb and 1.2 kB.both endhas  only  6bp  oberlaping region. The final product was successfully expressed and purified.
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Best in this case means: specific priming, long templates (amplifying whole plasmids >5 kb), and fast.
I use NEB Q5 polymerase now which meets all of these requirements, but is very expensive from my country's distributor. Q5 is a big improvement over what I've used previously (PfuUltra being 2nd best).
Q5 PCRs give good yield ~95% of the time on the first try and ~100% of the other times can be easily fixed with small tweaks, even with very poor primers (very low GC; many mismatches). A very good fidelity with 2 errors observed over ~1M bp of sequenced clones made from Q5 PCR products.
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Zach, Dima and I have been using KOD Neo Plus from Toyobo.  I haven't amplified anything over 3-4kb but he uses it routinely for plasmids and swears by it  Not sure the costs outside of Japan but it was definitely cheaper compared to other hi-fi long range polymerases. Compared to Q5 which I also use and feel the same way as you about it, it does better with less template.  I use Q5 only b/c they make a lazy man's version of it as a 2X master mix and of course, funds are not an issue in our lab.
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I'm trying to directly assemble oligo with flanking sequence into the lentivirus.
This method has been applied in CRISP-seq. They design the oligo(ssDNA) with NNNNNNNN and flanking sequence and use Gibson Assembly to gain plasmid pool.
But I can't figure out how oligo(ssDNA) could be assebmle into the vector? 
I see the positive clones myself. But there are still many false positive clones without oligo. I need at least 1000 clones with different barcode oligo. So I wonder whether the efficient of stbl3 could provide me enough different positive clones(>1000)? I used Single enzyme digestion because the oligo is too short(about 100bp).
Should I improve the purity of my cut vector in the assemble reaction? I think the false positive could not be totally avoid in Gibson Assembly with single enzyme digestion, I could make a higher efficiencies in assembley, but I am not sure the problem is the purity of my vector or the ratio in the reaction.
Thank you!
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Yes this is certainly carryover of uncut vector.  Gel-purification should be adequate.  But even better, make the vector by PCR from a very dilute template.  (as low as the PCR will allow).
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I am doing Gibson Assembly (overlap pcr cloning) and I am trying to clone a gene into pET22b directly behind the pelB leader sequence. I'm facing a problem with high GC content in a few of my primers, at the moment the smallest percentage i can reduce it to is around 80% GC content. Does anyone know if this is possible to amplify and if it is how can i improve the fidelity of amplification during pcr?
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Add 5-10% DMSO in PCR system and do a gradient  PCR, the annealing temperature accross 60-65 ℃
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I have used Taq as my enzyme...
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I am not sure about what type of PCR template you have got.  I think Taq Pol. may not be a good one (if the template is large, such as a vector), and DpnI digestion will help you degrade the original strand. If you have not checked already, you may benefit from the following link:
All the best with your PCR mutagenesis!
Best, 
Mahi
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Hi everyone,
I'm trying to perform an assembly using the NEB Gibson assembly kit.
My aim is to assemble 3 fragments : one of 421 bp fragment {my gene of interest} , and 30bp{the fragment  can make my protein accumulate in vacuoles}, and my vector backbone of 2376 bp. 30bp is so short, I cannot use NEBuilder Assemble tool to design primer.
Or which ways can I use to ligate the 30bp to 421bp fragment?
Forgive my English, I don't know  if I explain question clearly.
Can you help me with it please, maybe you have some good advices?
Thanks a lot!
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You can make it artificially longer, by overlapping it with vector sequences.  Just order a gBlock for a 500 bp of vector sequences containing your insert, and assemble that into an appropriately shortened vector PCR product.  You're sequence-independent now!  No more constrains on your imagination :-) 
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I have three gene segments (A,A,B)- note that two of the fragments are identical in sequence. I would like to prepare the following three constructs
AAB, ABA, BAA in pcdna3.0. 
I would like to stick with a seamless cloning approach so Gibson came to mind. For those of you that use Gibson, do you see a potential problem by the repeat sequences? If there is a better alternative I am open ears. 
Best,
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Hi recently I did a Gibson where the backbone contains already a gene A and I wish to add additional two A as the two inserts, so the target plasmid should have AAA.
The backbone is cut by one restriction enzyme MluI, and I carefully check if the three overlapping regions between backbone-insert1, insert1-insert2 and insert2-backbone are independent.
However, I could never achieve this goal by picking up in total 16 colonies and I never met this ratio of negative result from Gibson cloning with 2 inserts.
So now I decide to turn up to using traditional enzyme cloning and insert the two inserts one by one. Although takes more days, but from my experiences with more successful ratio.
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I attempted gibson assembly cloning on my gene of interest and the PCR product was correct (attached image). The transformation (DH5-alpha cells) was successful, but when I ran an analytical digest, a band size of ~450 bp showed up when I expected a 328 bp fragment (according to my SnapGene map). 
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My guess would be that there's something happening during the assembly process that makes it possible for multiple copies of the same insert to fuse together into one piece. Are there large(ish) repeat areas within your sequence or anything like that? 
I've done a few Gibson assemblies using a tandem dimer protein, and it always gives me weird products, like apparently monomeric and tetrameric insertions of the protein. The only thing that seems to work is just to screen as many colonies as I can until I find one that's correct. Another option might be to change the fragment ratios, as that might make the reaction favor the product you want. 
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I'm attempting to clone a gene into a vector via Gibson Assembly. My PCR insert band is the correct size (~1500BP), I did the Gibson assembly reaction and transformed the product into bacteria, did a miniprep, and digested at the restriction sites that i placed my gene into the vector. For some reason I have a band that popped out WAY bigger than my vector in two out of three of my preps
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Hi Eric, the bands you are seeing both derive from the same plasmid DNA. The faster band is supercoiled, the slower band running higher up in the gel is nicked/relaxed, but still circular. If you linearize the plasmid with a single cut, you should see a band that migrates a little more slowly than the supercoiled and faster than the relaxed circular DNA.
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I'm having a lot of troubles with this reaction. Supposedly, (http://www.synbio.org.uk/dna-assembly/guidetogibsonassembly.html) it's pretty quick and easy... but I've been working in this construction for months. I'm working with an autoreplicative cDNA, 5 of my fragments coming from RT-PCR, and all fragments are very variable in length (0.250 - 4 Kpb). I purified all the fragments by GE columns and quantify by nanodrop and gel visualization. I don't know what else I could change...
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Gabriela, I never tried to fuse 15 kbs, but I did 6 kbs succesfully. Maybe you can try assembly the smallest fragments with overlap PCR, and then try the gibson method... Use a good high-fidelity polymerase. Sorry I can't help you more... Good luck!
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Ligation reaction approximately 5000 bp.
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How do you check you ligation? via Agarosegel electrophoresis? Could it be that your insert is toxic or something like that? Otherwise you may try it again with other chemocompetent E. coli from your lab. I used to use DH5alpha and it worked very well together with Gibson assembled constructs.
Did you use the positive control given in the kit? If that works maybe something is wrong with your assembled product or the competen cells from the kit and you will be able to check your method is not going wrong.
But just guessing what was spontaneous in my mind. Hope you will be able to fix he problem.
I'll keep my fingers crossed for you.
Sara
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We're working on cloning for C. elegans, and we're having difficulty with what I assume is plasmid uptake.
We're amplifying out a 6kb segment (vector + rgef promoter + RFP), and a 5.6kb (dip2cDNA + 3UTR + GFP) segment, then ligating them together using Gibson assembly.
After inoculating plates and incubating overnight, very few colonies have grown. and upon PCR screening for inserts, very weak results were obtained.
If the DH5a cells are the issue, are there any other non-methylating bacteria that are good for larger plasmids?
I suspect the size of the plasmid is affecting efficiency. Anybody have any suggestions?
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Hi Cynthia,
do you mind sharing your protocol for DNA purification by low-melt agarose and Beta-agarase digest. Then concentrate using a Millipore DNA Fast flow ultracel Microcon units?
Thank you very much!
Gang
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Dear All
I am going to set up Gibson Assembly reactions. The reaction takes place in isothermal buffer which requires 100mM NAD and I have not been able to find this concentration of NAD. I would appreciate if those of you who are already doing this experiment, would let me know where to get it from. NEB sells NAD but it is a 200uL vial with just 50mM concentration.
I would appreciate your help.
Thank you
Ikram
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If you want to save money but still use the commercial kit to avoid possibly complicated troubleshooting, know that you can greatly reduce the reaction volume that is stated in the commercial datasheet. NEB indicates reaction volumes of 20uL (thus 10uL of 2X gibson assembly master mix per reaction) and subsequent transformation of 2uL of this reaction into competent bacteria, but I scale it down to as little as 2-3uL total reaction volume (1.5uL of 2X mix) and transform the whole reaction into bacteria. This way, I can make at least 5 times more reactions with one kit.
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Dear All,
I am going to set up Gibson Assembly reactions in order to clone certain fragments in a vector. The reaction requires three main enzymes i.e. Phusion Polymerase, Taq Ligase and T5 exonuclease. My question is whether I would need all the enzymes from the same company or not? Do I require to supply specific buffers like buffer for Taq Ligase; specific buffer for exonuclease etc. If yes, then how should I establish my reaction? Also, I am not using any kit.
I would appreciate your help.
Thank you
Ikram
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Hello All,
I am currently struggling with getting a Gibson Assembly to work. I am trying to insert a 2.5 Kb insert into a roughly 12 Kb plasmid (modified pLEX-MCS). I have had the G.A work in the past with the same plasmid.
Currently, I am creating the insert with 25 BP of homology on both ends via PCR with Q5 polymerase, this is the same homology sequence I have used in the past. For the G.A itself I am using ~60 ng of the vector and ~40 ng of the vector which is 3:1 insert to vector. Incubating at 50 C for 60 minutes followed by transformation using 3uL of the product. Transformation of the G.A product, cut vector and insert result in no colonies while ~10ng of the uncut vector results in a field of bacteria on the plate.
Side note, the gene I am inserting is non-toxic to bacteria, I am PCRing the insert from a pBabe vector which does not seem to affect the bacteria containing it,
Thanks!
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Thank you for your suggestions. I went back to square one and re-purified my vector and insert and did a 4:1 insert to vector ratio and ended up with ~60 colonies. Using PCR I have found that 9 out of 10 colonies tested have my insert. I am going to chalk it up to older plasmid/ potentially degraded DNA. 
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I want buy this kit but I do not know anything about efficiency of this kit. this kit manufactured by NEW ENGLAND BIOLOB.
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I did try using the NEB kit and yes it is fine you can get colonies with the right fragments you want. But we tried making home made Gibson reaction mix and it works fine and same result as the commercial kit. The only advantage is you can save more money here. If you are interested I could share you our cocktail mix.
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I'm trying to express a fluorescent protein (mNeonGreen) from the SV40 promoter of pcDNA3.1. The plasmid was obtained via replacement of the BleoR gene with mNeon using Gibson assembly. When I transiently transfect into both HEK293 and HEK293T, I see no fluorescence with the green filter. Our lab has another pcDNA3.1 variant in which the CMV promoter is replaced with the Ubiquitin promoter; I use this weaker promoter to drive RFP as a transfection control. I co-transfected this control plasmid at 1/10 concentration of the GFP plasmid and observed robust red fluorescence in the same plates. Additionally, replacing the SV40 promoter with a CMV promoter resulted in robust green fluorescence, suggesting that the problem is with SV40.
Sequencing suggested everything correct in both the promoter and the gene, save for a single nucleotide deletion at the 5' end of the promoter (image attached). I downloaded the sequence of Invitrogen's pTracer-SV40 expression vector and it's also missing this nucleotide so I don't believe it's crucial. Again, if I do nothing but swap the SV40 promoter for the CMV promoter, I get robust expression. What could be the reason for my expression failure with SV40?
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Dear all,
I was wondering if anyone has made any inroads in explaining this? I too am having trouble with my HEK cells. I have been attempting to transiently transfect HEK-293 and HEK-293T cells with a multi-part plasmid construct, that expresses a fluorescent reporter (Fast-FT) from an SV40 promoter. I have been getting no fluorescence, and the controls (using a CMV promoter) have been working. When repeating in CHO, it seems to work (although with low efficiency).
Vu Anh Truong were you able to find any support for your silencing idea?
Interestingly, ours too seems to have a point mutation. But since it works in CHO, I am hesitant to believe that this is the issue.
Thanks,
Alexandra
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I am looking for pTRIPZ vector with a resistance other than puro.
I would appreciate if someone that built it already could share.
I tried to replace PuroR by HygroR using Gibson assembly but my PCR for the vector (with PuroR deleted) keeps failing using Fusion or Accuprime (12kb).
I attach the map and would appreciate any advice on an alternative cloning strategy.
Thanks,
Igor
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I haven't worked with TRIPZ directly, but in analogous cases I have been successful using QuikChange from Agilent.
I would first remove the pac gene using QC "deletion" protocol, and then insert the whole hygro (or similar) resistance gene using QC "insertion" protocol. I have used the QC kit to introduce fragments up to 1.2kb, with good results.
Good luck.
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I’m trying to assemble 4 fragments (2.1kb vector + 3 inserts, each around 1.1kb) using Gibson assembly. All three inserts have the same sequence at the beginning and at the end (200bp promoter and 200bp terminator). So I added custom overlaps, each between 30-40bp to ensure the proper order of the assembled fragments.
The problem is that I’m not able to obtain colonies with the correct insert (= vector with all the 3 inserts in the proper order). I’m getting colonies with 1 or 2 inserts, sometimes even more, but it’s a random mix of insert fragments in the vector.
After the assembly, I can see very strong bands that correspond to each of the fragments, and weak bands for combinations of 2 fragments, but no bands for 3 or 4 fragments together.
I’ve optimized the overhangs in order to remove all potential secondary structures and also the GC content is below 50%. Using the same overlaps, I’m able to join each of the insert fragments into different vectors.
The Tm temperature of the overhangs is 60C, except for the first insert, where the Tm is 67C. Do you have experience with the Tm of the overhangs? NEB recommends Tm > 48C, overhangs in the example in their manual have Tm=50C, so maybe my Tm is very high for assembly of more fragments than 2? Incubation time is 1h at 50C.
Thanks!
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Yes I think more than equimolar inserts is an excellent idea too.  Try that first, it might do the trick. 
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I'm trying to insert a 120bp segment into a 4k vector using Gibson Assembly. The purpose is to replace a 27bp portion of the vector with NNKs. The insert is ordered as an ssDNA oligo, and then made double-stranded after annealing to a primer, and completing the second strand with Klenow. It has 36 bp overlap with the vector. The vector backbone is prepared using inverse PCR, because there are no restriction sites around the target area. I then use DpnI to digest the WT vector, and then gel extraction to remove all the DpnI fragments (which can be up to 1kb and tend to ligate back into the vector during Gibson Assembly, so gel extraction is necessary). After cleaning/concentrating the vector product with the Zymo kit, I mix the dsDNA insert with the vector backbone (1:5 vector to insert ratio) with Gibson Assembly, and then chemically transform into homemade MegaX (DH10B) competent cells. The goal is to maximize transformation efficiency, since I'm creating a library.
This has worked for me once, several months ago, using a similar insert that was only 87 bp long (21/18 bp overlaps). Since then, however, it has consistently failed-- I get zero colonies, or at most, a few tiny colonies. I later switched to a larger insert (120bp) since some said that the T5 exonuclease of Gibson Assembly could totally chew up a small fragment. Still, no luck! I can transform the original vector template successfully each time, proving that the vector, antibiotic resistance, transformation protocol, and cells are all good. I've also run a positive control with my Gibson mix, which has worked.
I get colonies if I skip the gel extraction step, but then half of the colonies contain weird combinations of the insert and DpnI fragments that were not removed, so that's not desirable.
Any other thoughts for troubleshooting? I can't understand why this worked over several repeats in the past, and suddenly cannot work anymore. Thank you!
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Hey Shivani,
I had a very similar problem with no colonies when I used gel extraction. Turned out that exposing the gel to uv light was the problem. I did control experiments and found even 30s on the lowest setting reduced transformation efficiency ~100x. I would recommend using sybersafe to stain the gel and cutting it on a blue light. Hope that helps.
Alex
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I am trying to put together the promoter, gene and terminator in pUC19 backbone. I followed the protocol of NEB in the Gibson mix. But  I am not getting any colonies in my plate ( even blue colonies). I made adjustment already in antibiotic concentration, volume of Gibson mix in transformation and even molar ration of fragments and backbone yet still not working. Can you suggest any information or protocol for the Gibson experiment?
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Since you don't have any background colonies,
I would suggest that transformation is not working.
Check your competent cells, if possible, switch to electroporation.
 
I would not worry about the [Amp] since bla resistance marker
can allow bacteria to grow on 30 times higher [Amp] than the
commonly used (100ug/ml).
Instead of the GIbson assembly, you may try to assemble your
insert from fragments by running 5-7 cycles of high fidelity
PCR  - larger product formation should be visible on the gel.
 
 
 
 
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Dear All
How do I decide upon the proportions for Gibson Assembly. I have a linearized (digested) vector 2958 bps (27 ng/ul) in which I want to clone two DNA fragments. One fragment is small 410 bps (80.1 ng/ul) and another is big 2750 bps (66.2 ng/ul). I have a 15 ul 1.33X master mix for assembly and now would need 5ul of the vector+small fragment+big fragment. How should I decide upon the proportions.
I would appreciate your help.
Thank you
Ikram 
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Hi Ikram,
equimolar amounts for all fragments are recommended. For example if you use 100 ng of the vector (3.7 µl), this would be 14 ng (0.2 µl) for the small and 93 ng (1.4 µl) for the big fragment. But this mixture would exceed the maximum volume of 5 µl, so just scale down the amounts to fit to 5 µl. If the assembly won't work using these amounts of DNA, I recommend to scale up the amount of DNA input for the small fragment (try 2 times and 5 times higher amount).
Good luck
Gertrud
Is anyone else Using gBlocks and Gibson Assembly to make phage display libraries?
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I'm a newcomer to the field of phage display (but not molecular biology or biochemistry) and I was wondering if I'm the only one making libraries from gblocks (containing randomized nucleotides at certain positions) using gibson assembly. It seems much easier than kunkel mutagenesis, or some of the other expensive nucleotide engineering approaches. Is there a downside to this strategy that I'm not seeing?
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seamless is a good choice for constructing synthetic antibody/peptide library. I ever constructed a lib in this way. for immune or naive lib, we used the classical method.
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I use Gibson Assembly routinely but find that about 25% of the clones I generate have errors in them at or near the site of assembly. I use the NED Gibson Assembly master mix but reduce my reaction volume to 4 or 5 uL. I talked to NEB about this and they suggested that this error rate is expected. I'm having trouble believing this because if you're stitching together 4 or more fragments this error rate would almost make it impossible to get an error free clone. Does anyone else have this trouble? Can this error rate be due to reducing the reaction volume? Do you think that the original Gibson Assembly mix would perform better than the one NEB makes?
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I am now using the high fidelity mix and I seem to be having much better results, although I never tried both mixes head-to-head on the same targets.
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Gibson assembly is supposed to be seamless in cloning especially when you want to make a construct from different pieces (more than 2). Since the commercial kit from NEB is expensive, I would like to have my own home made kit.
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I was wondering if anyone had experience with using Gibson assembly for stitching Oligos. I've never tried it before and I'm willing to give it a go. (i.e. using a small oligo which overlaps the vector and the insert as a stiching oligo).
I have a vector which is 7kb long and an insert 2kb long. Both were digested with restriction enzymes.
Does anyone have a tried and tested protocol?
Otherwise some specific questions I thought of included;
i) how long an oligo? I'm using 60bp, 30 from the insert, 30 from the vector
ii) did you need to phosphorylate the oligo?
iii) how much oligo to use?
iv) % success in your experiments.
Any other tips from experience would be much appreciated.
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See figure 1 here for the mechanism:
The 3' homologous sequences of the vector and inserts are reversed. Isn't this a problem?
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No, they are not. The primer is of course the reverse complement, but in the final plasmid the sequence is in the correct order. The Figure might be a bit misleading, but try to draw it yourself with an actual sequence and that become apparent. If necessary, I can send you a better figure (I think) early next week.
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I am trying to generate a 15.3 kb vector. This is split into the vector backbone (2.1 kb) and a large piece of DNA (13.2 KB) which I have split across 3 PCR fragments (6.2 KB, 3 KB, 4 KB) – thus I am trying to assemble 4 fragments. I’ve ran all the fragments out on a gel and they look fine. The DNA concentrations are between 16-100ng/ul. There is minimum 20 bp overlap between fragments. In the Gibson assembly reaction I’m using equimolar ratios, (calculating from 70 ng of the vector) and incubating at 50 degrees for 1 h. My total reaction volume is 40ul (20ul DNA and 20ul mastermix) and I heat shock transform 2 ul of this using the NEB 10-beta cells. I tried several times, and there are colonies on the plate yet I never get the right product after testing all colonies. The positive control with the kit works fine. I was thinking that perhaps the overlap isn’t long enough and considered doing the reaction in 2 stages, 1st incubate 3 fragments with the mastermix for one hour, then add the third fragment (6.2 kb). Any suggestions of the best way to try this as I’m now quite limited on the amount of mastermix I have left?
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I'm guessing this most likely sounds like a secondary structure problem. Gibson Assembly is not exactly as sequence-independent as the headlines would have you believe. You need to avoid G/C rich regions like the plaque. If it has as little as 5 GCs in a row in your overlaps, or an overall G/C content >65%, that alone can ruin a single reaction. And when you do 4 of them at the same time, then those probabilities compound against you.
Longer-range structures can be a problem too. If you assemble innocuous sequences that work in another context, the same assembly does not necessarily work when it's near structure-intensive features like IRES, WPRE or Lentivector LTRs. I've used secondary structure prediction software tools with some success to call this problem in advance.
Length of the overlap doesn't really matter (except as a function of Tm and G/C). The Tm needs to be in the 45-50C range. There is no advantage to making it longer. Again, you *must* resist the temptation to increase your Tm by operating in G/C rich regions. You will save yourself a lot of trouble if you have the freedom to operate near balanced to low-GC regions and away from structures.
If it isn't obvious from the above which junction causes your problem, then yes, do break the reaction down into single-step assemblies as much as you can. That will improve your efficiency. And you need all the efficiency you can get for this type of project.
You can conserve Gibson reagent by reducing the reaction volume. There is no reason to electroporate 2 ul and throw 38 ul away. You can totally scale everything down to 4 ul or I've done even 2 ul total volume. You're going to be limited by evaporation of reagent during the reaction. Pick the smallest tube you can get, and immerse the whole vial in a waterbath, to eliminate small differences in temperature on the tube walls. Just make sure you get the math right. All the concentrations need to stay the same, while the volumes and absolute amounts all go down. Excel is going to be better at this than your brain ;-)
The second method to conserve costs is to buy the components and mix them yourself. It's unbelievable how much margin people can make for filling things into single little vials for us!
Keep it up though. If you have to clone a lot, then mastering the idiosyncrasies of this method is going to be extremely rewarding in the medium term.
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I used 100ng vector. I think the Gibson assembly is low efficiency to get ligation.
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Hi Liang,
I typically use 25 ng vector and a 3:1 molar ratio of insert(s) to vector. I normally prepare vector by digestion and inserts by PCR. I make vector and insert up in 2.5 µl volume and add 2.5 µl of the 2x master mix (from NEB) and then do a 60 min reactions at 50°C. If you are using NEB competent cells, then adding 1 µl of this reaction to 20 µl of cells and doing a transformation, adding 100 µl of SOC for recovery and then plating *everything* typically gives me 100-200 colonies on a plate.
So, it may be that, counterintuitively, you are using too much DNA. I would recommend using less DNA. It is also important not to add too much volume of transformation mix to the competent cells - 1 µl in 20 µl or if you are getting no colonies maybe 2 µl in 50 µl.
The manual also states: " Use the competent cells provided with the kit (NEB 5-alpha, NEB #C2987). The components of the Gibson Master Mix may inhibit the functionality of competent cells from other companies." If using cells from other companies you may need to dilute the reaction mix 1/4 and then use that to do the transformation.
Hope that helps.
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I started using Gibson Assembly from NEB in May, and I got fantastic results. Using the simple NEBuilder, and the chemically competent cells in the kit, I would get a lawn of colonies, every time. I returned to cloning this September, bought a new kit, and its been a disaster. Using the NEB chemically competent DH5 alpha, my positive control would only give me a few colonies, and my pUC19 would work even worse (transformation efficiency 10,000 per microgram).
NEB has been great customer service wise, sending my two other kits, but again, my pUC19 efficiency is so low. They then sent me NEB 10 Beta chemically competent cells, and I'm not getting any colonies at all with it, just using the pUC19. For my own sanity, I just did a transformation with a plasmid I know that works and the pUC19 provided into chemcially competent cells I made myself, and my plasmid gave me a lawn (although it was at much higher mass) and the pUC19 gave me nothing. I've also tried using a plasmid I know that works into the NEB 5 alpha cells, and I get more colonies, but still, really low transformation efficiency. I've tried different plates, doesn't matter. The positive control is a little better, but still really low efficiency.
This is all so frustrating, because four months ago, I was getting tons and tons of colonies. I am about to try electrocompetent top ten cells and desalt my ligation reaction with drop dialysis. Does anyone have any other ideas? Is anyone else going through/ has gone through something like this?
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Are you cloning the same gene? It could be that the gene product you are cloning is toxic to the cell. Could you provide more details about your insert?
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I have been trying to assemble three fragments with sizes 2400bp, 770bp, and 1000bp with Gibson assembly master mix. The overlap between individual fragments is 45 to 50 bp. The GC content in the overlapping regions is quite OK. I followed the recommendations given by NEB for the assembly reaction and a few variant conditions without success.
Is it fine to store the Gibson assembly master mix at -80C instead of -20C (Recommended storage temperature by NEB). I don't think it is a problem but would be nice to know if somebody has a similar experience. Before facing the aforementioned problem i could successfully assemble three different fragments in the similar size range (2100bp, 770bp, and 460bp) using the same kit (The Gibson assembly master mix was stored at -20C at that time). Please share your opinion.
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Thank you Paul Jaschke for the suggestions. I shall try the two piece combination as you suggested. I tried assembly times of one hour or more, i think 15 minutes is too short, is'nt it?
I am not using the vector in assembly as such, what i am trying to do is assemble the PCR generated fragments initially and then blunt end cloning of assembled fragments into vector of choice. Please let me know if you have any suggestions in this regard.
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We aim to assemble 6 PCR fragments (300-400 bp) by Gibson assembly. We assembled and PCR amplified the first 3 and last 3 fragments with no problems. However, the assembly of the two amplicons to the full-length product fails and PCR analysis shows that fragment 5 and 6 are faulty. We are using the NEB Gibson assembly master mix essentially according to protocol. We assume that secondary structure during assembly of the two amplicons at 50 degrees C are causing the problem. Do you have any suggestions on how to solve this problem?
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I completely agree Karl and Abdelhalim. DMS0 (as suggested by gonzalo could be useful too).
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I do most of my pcr reactions in 10uL volume. I use Takara SpeedStar polymerase and a master mix made by someone else in the lab that includes buffers and all the necessary PCR things. My 10uL reactions get 1ng DNA, and 1uM primers (final concentration). For 3 different reactions, I got very bright, clean bands with no smearing. I needed to increase the volume to get a larger yield for gibson assembly. I kept all the final concentrations the same (i.e. .1ng/uL DNA and 1uM primers), but increased the volume to 50uL. My pcr machine has a setting for reaction volume, which I raised to 50. This time, I only got product in one of my reactions, and it was not clean. It was a huge smear. Halp!
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Can you check the cycle length? As your liquid will take longer to get to the correct temperature, there is more possibility for unspecific reactions or too short duration of correct reaction temperatures. It seems your machine aims to correct for this, but I would possibly check this and play around with cycle duration and speed of temperature increase.
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Good day to all. I am encountering a problem in determining the correct amount of plasmid and inserts' concentration. According to the Gibson assembly protocol, I should include a 0.2-1 pmol of vector for an assembly of 4-6 fragments, by using the formula appended below:
pmol = (weight in ng) X 1000 / (base pairs X 650 Daltons)
Also, the protocol suggested an optimum range of vector concentration of 50 ng-100 ng. Let's say if I were to use a vector of size 2000 bp, the pmol range obtained for 50 ng - 100 ng is 0.038 - 0.076 pmol, a range far from 0.2 - 1 pmol. Would appreciate if anyone could provide me suggestions and advice on this matter. Thank you.
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This doesn't need to be precise.  The technique is very tolerant to deviations.  I've done scores of different Gibson reactions and had all kinds of problems.  Not precisely calculating the masses was never one of them :)  It will generally either work really well, with most colonies being correct over a broad range of conditions;  Or it won't work at all no matter what you do. 
If insert and vector are approximately the same size, use 50 ng vector and 150 ng insert.  
If they are different sizes, correct by the ratio of their length, so that you add less mass of the smaller molecule (i.e. keeping the amount constant at 3x insert excess). 
If you don't get any correct colonies, it's not the fault of your precise DNA ratios.  It's the fault of picking GC rich overlaps that won't work, or some other problem.  
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can anyone please explain picomole calculation in gibson assembly like what is vector and insert ratio to be added in reaction and how to add  a picomole to a reaction?
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NEB recommends 50-100 ng vector and a 2-3 fold excess of insert.
You usually know the length of your vector and of your insert. Then you calculate or measure the amount of DNA in µg. The MW of a base pair is known (app. 660 daltons). Now you can start to calculate the pMol/µl of your DNAs. Example: In case of a 1kb fragment  0.66µg are 1pMol.
See the formulas in e.g. in the Roche "Lab FAQs"
or the calculation in the NEB ManualE5510 calculating with 650 daltons:
pmols = (weight in ng) x 1,000 / (base pairs x 650 daltons)
50 ng of 5000 bp dsDNA is about 0.015 pmols.
50 ng of 500 bp dsDNA is about 0.15 pmols.
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Hi all,
I am trying to assemble two fragments (insert and vector) with NEB gibson mastermix. I already have done several reactions using different insert:vector molar ratio (3:1 - 7:1). Several colonies formed from each reaction and there were no colonies in negative control plates. However, I could not find any inserts after either colony PCR or mini-prep. Are there any suggestions? Thank you.
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Sorry. I should have gone through my question before submission. Yes you are right,  insert:vector ~ 3:1 - 7:1. Overlapping regions are > 20 bp.
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After designing the primers for Gibson Reaction is there any free S/W to simulate the reaction and check? 
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On the NEB website you have an online tool to design your assembly and primers.
Check the link below
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I need to subclone a gene into an unusual vector that has only EcoRI at the insertion site.  I don't have a construct in which my gene is flanked by EcoRI sites, so I will have to PCR amplify it to add them no matter what.  My gene also has an internal EcoRI site.  So, instead of doing a partial digest followed by non-directional cloning, this seems like a great opportunity to try Gibson Assembly.  I was thinking that I could digest the vector with EcoRI and generate my insert by PCR with primers adding 30 bp of vector sequence on each side.  My question is, won't the vector anneal to itself and reclose at a high frequency?  Perhaps the reaction temperature will be too high for a small overlap to anneal and the insert will be favored?  I know the other approach is to amplify the entire vector to create a blunt insertion site, but I'm worried about introducing errors.  If anyone has any experience with this type of situation, I would appreciate any advice.  Thanks!
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It is not advisable to linearised plasmid by single RE, as the chance of reaneal is high. To prevent re-anealing, you should use 2 REs (which the ends are not complimentary) OR if you are using only single RE, dephosphorylate the plasmid with alkaline phosphatase (like CIP,  CIAP, TSAP, rSAP) to minimise self-annealing of the plasmid.
Blunt ends annealing is considered to have lower efficiency of annealing that sticky ends. Products from the non-directional ligation could be filter out easily with PCR (1 vector targeted plasmid + 1 insert targeted plasmid). You should have plenty of clones available for screening. 
With DNA sequencing analysis, you can confirm if your whole construct is correct. 
If you are only aim to insert a single fragment, and the overall final recombinant plasmid won't be too large, you may want to consider overlapping extension PCR (http://www.rf-cloning.org). This is easy to design and works well. You no need to consider about the RE sites. 
All you need is high fidelity polymerase with higher fidelity (like Fusion polymerase, Q5 polymerase, etc., not pfu). And also DpnI RE. The drawbacks of this technique are (1) you might need to screen 100-200 clones for some cases, (2) the amplification is linear, so the transformation rate is low too (works on lab prepare Cacl2 cells), (3) size of the overall plasmid should not be too large for the polymerase amplification (~10kb will still be acceptable). 
Good luck
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I am trying to assemble a plasmid for transfection by Gibson assembly. Because there are no unique restriction enzyme sites in the region I am trying to clone into, I am effectively trying to linearize my plasmid by PCR with two primers designed back to back (they were selected by the NEB gibson assembly program). I have tried conducting the PCR using both Herculase II Fusion polymerase (from Agilent) as well as the NEB Phusion high fidelity polymerase. In both instances, I get random bands that are shorter than the total size of my plasmid, suggesting that perhaps the primers are binding to another location in my plasmid, or there is a point where the enzyme may stop polymerizing.
Does anyone have any tips on how to check for these? or any suggestions for how to minimize unwanted bands?
The vector I am using was generated by TA cloning a previous PCR product into pGEMTez vector. 
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Hi
I had the same problem. I would recommend the following
1. If possible try to linearize the plasmid ( even if it cuts twice around the site where you want to insert your fragment). By doing this i could get my fragments amplified. Super coiled plasmid are notoriously hard to amplify irrespective of the size. Did you run your uncut plasmid in a gel to look for that?
2.I used Q5 polymerase from NEB, i could amplify 11kb fragments with ease.Its optimized to amplify GC rich areas among others with ease.
3. Use the NEB calculator for Q5 , as the cycling conditions are very different from the regular polymerase. Also make sure to only add the sequence complementary to your fragment of interest. I had put the whole primer sequence and i hardly got any product  as most likely there was little or no annealing of the primers. 
4. Typically when you have very little choice to insert your sequence at a specific point , those place are either very GC rich or have a lot of so called "Polymerase sink" (Pol sink is my terminology, not aware if that exits). I had one primer seq which had a bad 3' end stability and Poly X issues (use AMPLIFX or similar to check this), and just by changing couple of base pairs in either ends, i could get around that.
Hope this helps
BR
Joy
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Dear all,
I am currently working with thermophiles, and i am trying to transform one of them. I found a shutle vector which should be able to do so. This vector can replicate both in E.coli and in thermophiles, and has amp and kan resistantce.
I cloned my insert into this plasmid via the gibson assembly protocol (inserts : 350 + 400pb, plasmid : 6500pb). After transformation in E.coli, I have screened the positive clones by colony PCR, made liquid cultures and then purified plasmids. Unfortunaltely, I only got shorter plasmids, with some deletions I suppose. And of course the insert is gone. So apparently, from the colony on plate to liquid culture, something happen... All cultures are made with selection pressure (50ug/ml kan + 100ug/mL amp).
This plasmid has a AT of 60%, and I have read this could lead to unstability and deletions (due to promoter activity in E.coli of these high AT content regions).
I am know trying to grow the E.Coli cells at 25°C (both on plates and liquid cultures), since I have read it could decrease the unstability.
I am also considering to change the competent cells (currently working with NEB 10 beta). I may change for CopyCutter Epi400 (from Epicentre), since it decreases the copy number, and lead to better stability.
Does anyone have already try to clone high AT content insert and/or plasmid ? Or get trouble with stability? Or loose the insert somehow ?
Thanks in advance for your feedback and your help ! :)
Antoine
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I tried the Stable competent cells from NEB, and I got nice results. They have specific deletions, and you have to grow them at 30°C. I've made 8x5mL cultures from the transformation plate, followed by miniprep. I got nice plasmid concentrations (all around 400ng/uL), and they are all pure, without degradation.
Thanks for you help !
Any advice with an Issue with transformation after Gibson assembly?
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Dear All I used the NEB HiFi builder to assemble my plasmid. The vector is 8.5kb, Insert 1 = 4.5Kb and insert 2 =1.2kb. I get my fragments assembled and can visualize in agarose gel. When i do the transformation, i don't get any clones. I have previously used the Mix & Go (Zymo research) reagent to make competent cells and it work very fine even with gibson. I tried using Stbl2,Stbl3, Endura and DH5A-Max efficieny-t1 cells but with no success. I have tried diluting my reaction mix 4 fold and used 2 to 5% of the diluted or undiluted reaction mix for transformation. Still doesn't help.  Can anyone please help
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I had issues the first time I used Stbl3 with the NEB HiFi builder. It worked perfectly when I tried the NEB stable bacteria.
Any suggestions on my transformation problem, with many colonies but not positive ?
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I was trying to clone 2kb gene into pet17b vector using both gibson assembly method and conventional method. After transformation into DH5alpha cell, i used to get 30-40 colonies and all were negative. I sequenced my pcr product and observed that it was correct. Can anyone help?
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Hello Archana, Check the quality of your starting linearized pet17b vector. How did you open it? Have you taken dispositions to minimize its recircularization ? The ligation of the two extremities of the linearized plasmid is an intramolecular chemcial rpocess, whereas the ligation of the open vector and an insert is an intermolecular process. Intramolecular processes are much more probable than intermolecular ones. If your linearized plasmid is prone to recircularization (due to a bad dephosphorylation for instance), your results are quiet normal unfortunately. Best regards Philippe Urban
Failure with Gibson assembly and CPEC cloning building 10kb plasmid?
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I'm trying to build a plasmid which is 9100bp size. I used Gibson assembly and CPEC cloning to join 3 fragments to build this plasmid. I got a bunch of colonies after transformation. I screened about 20 colonies and all of them have truncation (a big piece of plasmid is lost). I sequenced the truncated plasmid and it shows that the junctions between 2 fragments are still there. It looks like the lost portion of the plasmid is recombinated to the genome or somewhere else. Does anyone had this experience before? How to solve this problem? 
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I had a similar problem. Each PCR fragment was perfect before the Gibson assembly, I could transform and select many clones. When I tried to do a PCR to amplify this plasmid I noticed that something stranged happenned. I solved this problem: in my case, I was trying to assemble the p15A origin of replication with the ampicillin gene resistance and a few other parts to build a new custom plasmid. All the sequences I used were from the BioBricks repository. I noticed that the p15A fragment I was using was carrying a region of 250pb which was identical to the end of the ampicillin gene resistance. During the Gibson Assembly, the two identical regions recombined (they were pretty close to each other) in a strange way and a big part of my plasmid was lost. I had to remove the unwanted homology region to be able to build my plasmid and then everything worked perfectly. Thus I will advice that you check carefully the sequences of each part you are trying to assemble together. I hope you will fix this problem soon!
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I'm designing an assembly and plan to use IDT gBlocks to generate my insert sequence. Using the gBlocks I can add the reverse complement of my vector primers to the ends of my gene insert, effectively leading to 100% overlap between 18-24 nt at the ends of my vector and insert.
When I analyze this plan in the NEBuilder Assembly the output flags potential mis-annealing. I'm confused as to why this would occur as long as I complete a Dpn1 digest to get rid of background vector. 
Does anyone have a recommendation for primer design in this case? (I've already checked out Janet Matsen's and the Miller lab's guides)
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The sequences you add to the gBlock shouldn't be "reverse complement" to the vector overlaps.  They should be identical to the vector overlaps.  For example:
vector-AAAGA         AAAGA-yourgene-CACCC         CACCC-vector
-> Both overlaps are identical to the vector sequence, not reversed or complemented.  You make the vector by regular PCR, using forward primer CACCC and reverse-complement primer TCTTT.  (If this didn't make sense, try to draw it out double stranded on a white board and follow the mechanisms along for both PCR and assembly.  It helps to see that.)
Technically you don't need NEBuilder's opinion on this.  If you followed this scheme, the IDT-calculated Tm of the overlap is >48, and your GC% isn't crazy, it will work. 
DPN1 usually not necessary, if you make the vector by PCR.  Just dilute the template plasmid 1:100.  PCR will still work fine, and background colonies are so dilute you won't ever see one. 
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Hi, I am designing a cloning to produce 2 different fusion proteins. The maps would be:
1- Gene1(1kb)--GeneX(2kb)
2- Gene2(7kb)--GeneX(2kb)
My question basically is, which are the principles I should follow in order to design the linker?
P.D. If it is important at all I am going to use Gibson assembly.
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Hi Inigio,
Can you describe what you are trying to achieve with the linker exactly? There are standard linkers that are used but may not be appropriate in your case. For example:
The assembly question is secondary to your design question. Once you know what you want to build you can figure out how to build it. Gibson assembly is a very useful method along with many other similar methods that rely on homologous recombination. These methods suffer when the DNA you are assembling has repeats or very high or low GC-content. If your proteins and/or linkers have these features you may want to use a different method to assemble such as restriction enzymes.
I hope that helps!
Paul
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Me and a postdoc in the lab are in dispute over this. I say that a 213bp overlap is too long because each end has to be chewed back by 100+ nucleotides before the insert and vector begin to anneal (thats why we have no clones). He disagrees. Any mediators?
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Thanks Di Xia for your input.
I actually did a test thinking along the same path as you just suggested. What I did was run the sample at 37C for 10 min prior to the 50C 30min incubation. My idea was that the 37C would give the T5 extra time at its optimal temp to chew back more of the overlaps (as opposed to adding more T5 which would probably have worked as well, like the paper said). And indeed, it looks like a success! I have now this morning 500+ clones (as opposed to 5-10 that I had before without the 37C incubation which were all wrong). I will check by sequencing to verify of course, but I am optimistic.
Thanks again for the help!
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I am using the NEB HiFi DNA Assembly Master Mix to assemble 4 fragments (about 1000bp for each) to pUC19(2700bp). The total length is about 7500bp. But I tried several times, I didn't get any colonies. My ratio is 1:1:1:1:1. I have checked my overlaps and the length of overlap is 35-65bp and Tm is about 70 degree and GC content is 40-60%. Can anyone give me some advice about my questions. Thank you.
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No colonies doesn't mean the assembly failed.  It means your transformation procedure isn't good enough to see it.  
If your transformation was up to snuff, you would be seeing false assemblies and vector background colonies (even when, and especially when the assembly fails!).  If the assembly works, then you get additional colonies containing the correct product on top of that.  It's virtually impossible to get zero colonies with a commercial, well-characterized cell that's certified for large plasmids. 
Your correct product might well be there.  If you can fix your transformation, you can find out. 
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Hi,
I am trying to insert a ~0.5 Kb fragment into a ~8 Kb vector by Gibson assembly, with no success (no colonies!).
Unfortunately the homology overlaps (20 bp long) have ~65% GC and each a small hairpin, which I think added to the size of the vector might be the problem.
I am quite constrained about these two features, I am afraid.
Is there a workaround this? Would it help to increase the length of the overlap?
Thanks
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UPDATE
Thanks everyone for your recommendations.
I tried again doing a transformation control in parallel, using the same vector I am trying to assemble in my insert. I almost got a confluent plate with ~40ng of a 8Kb vector, which is not a very stringent test but it passed. I am using Invitrogen one shot TOP10 cells anyway, so this was just to make sure they have not gone bad. I would deduce from here that there is nothing wrong with the transformation - it is just that the double enzyme digest and alkaline phosphatase treatment is efficient at preventing re-linearization.
This time I also made the assembly with more insert, which I had purified without running it into a gel (DpnI degradation of the plasmid template) - I read somewhere this can be important.
I think I am going to try next longer homology stretches - but please do let me know if you have alternative suggestions!
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As mentioned above.. But consider making competent cells using CMMB80 buffer method http://openwetware.org/wiki/TOP10_chemically_competent_cells (you can use another strain)
this should give cells with good enough competence! The competence of cells is critical...20 basepair overlap should work just fine.. As a control same amount of DNA with just water (= not Gibson Assembly master mix). Of the Gibson Assembly mix, don't clean up.. And use 5µL to transform 100µL competent cells. 
You can also increase the ratio vector:insert = 1:10, in 20µL with about 25-100 ng of vector.
Cheers
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I am using the NEB Gibson Assembly kit and failed to get any colonies for 8 different type of constructs, a several times, my control plate has only 25 colonies using NEB 5-alpha competent cells.My homologies are 22bp long.But my main question here is how many colonies should be there on the control plate in general.Please contribute as to what do you think the reason could be?
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Hi there,
Do you perform transformation with the full control reaction mix? If it's the case dilution  of the mix prior transformation may improve transformation efficiency.
A remark for John: if you don't get any clone from transformations using Gibson assembly reactions, I agree there might be a transformation problem as you said but there might also be a problem with the assembly reactions. To have a clearer picture, a transformation control with a circular plasmid should be done in parallele to check bacteria competence.
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I am using pTD-NStrep-His vector. I have cloned two genes in two individual vectors. Those genes are two cellulase enzymes. I used Gibson kit during the cloning process and all results were great starting from amplification of gene to cloning into E.coli BL21 (DE3). 
I have added both His-tagX6 and Stop codon (TAA) to the reverse primers so i don't have to add extra amino acids to reach the histag from the backbone vector.
The total concentration of Streptomycin (25ug/ml). 
The expression was carried out at 37C until OD 0.7, Induction with IPTG was at final concentration of 400 µM. Samples were incubated at 25C for overnight. Then cells were harvested and extracted using 1% Triton X-100 + 1 M urea.
By checking on SDS-PAGE, i could not detect any of the proteins in both cultures. Those are the SDS-PAGE of the purified histag and the elution fractions for both samples.
The enzymes MW are 35 and 44 kDa.
Histag1: the first enzyme fraction bonded to histag
Elution1: the eluted fraction (un-bonded) fraction
and so on..
As i ve gone through before, this enzyme has been produced by few researchers without using signal peptide. Thus, enzyme will be active by extracting from the E.coli cells.
The frame was sequenced and ligated in the proper frame.
However, I am wondering if i am facing one of these problems.
1- The vector pTD-NStrepHis is an expression vector which works by restriction enzymes as mentioned in Addgene database (supplier).
Is it ok to use Gibson assembly kit in this case ?
2- The enzyme could be produced as inclusion bodies and precipitated? If so, how can i analyze/extract the inclusion bodies for SDS-PAGE? 
Any advise? or other problems i might have ?
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Hi there,
There is no simple answer to your question... To have a clear picture of the situation, you should compare SDS/PAGE patterns of samples obtained with the same amount of non transformed cells, non induced transformed cells and induced transformed cells. It's hard to state on expression/no expression if you don't have any referential to rely on (especially if the expression level is quite low). In a second move you would have to state on the solubility of the protein expressed by comparing SDS/PAGE patterns of total crude lysate and supernatant of centrifuged crude lysate. If your proteins are expressed in inclusion bodies you should spot them in samples from total lysates or cells and get nothing in lysate supernatant.
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I am using pTD-NStrep-His vector. I have cloned two genes in two individual vectors. Those genes are two cellulase enzymes. I used Gibson kit during the cloning process and all results were great starting from amplification of gene to cloning into E.coli BL21 (DE3). 
I have added both His-tagX6 and Stop codon (TAA) to the reverse primers so i don't have to add extra amino acids to reach the histag from the backbone vector.
The total concentration of Streptomycin (25ug/ml). 
The expression was carried out at 37C until OD 0.7, Induction with IPTG was at final concentration of 400 µM. Samples were incubated at 25C for overnight. Then cells were harvested and extracted using 1% Triton X-100 + 1 M urea.
By checking on SDS-PAGE, i could not detect any of the proteins in both cultures. Those are the SDS-PAGE of the purified histag and the elution fractions for both samples.
The enzymes MW are 35 and 44 kDa.
Histag1: the first enzyme fraction bonded to histag
Elution1: the eluted fraction (un-bonded) fraction
and so on..
As i ve gone through before, this enzyme has been produced by few researchers without using signal peptide. This is an example (attached).
However, I am wondering if i am facing one of these problems.
1- The vector pTD-NStrepHis is an expression vector which works by restriction enzymes as mentioned in Addgene database (supplier).
Is it ok to use Gibson assembly kit in this case ?
2- The enzyme could be produced as inclusion bodies and precipitated?
3- The reading frame might be ligated wrongly, which is not common using Gibson assembly, so i sent sample for sequencing. 
Any suggestions ?
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I'm sorry to say that but that's really annoying!
You haven't even sequenced the your construct and are trying to get a protein expressed without knowing if your sequence is correct??
Sequencing is not an optional step in working with DNA, it is THE first and most important thing BEFORE you start doing anything with that construct! If you thought you would save time or money guess how much of both you have wasted now.
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Dear All
How many nucleotides does the 5' exonuclease remove during Gibson Assembly. I mean I was wandering how does the exonuclease recognize the position where it has to stop removing the nucleotides.
I would appreciate your response.
Thank You
Ikram
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Hi Ikram,
The number of bases the exonuclease removes is not a specified or determined number in a Gibson assembly reaction. This is a dynamic reaction that is "balanced" by the polymerase activity which is also present in the reaction. Think of these two enzymes constantly removing (exonuclease) and adding (polymerase) bases. Every now and then (hopefully more often than not!) the exonuclease will remove bases and allow 2 of your fragments to anneal, thus allowing the polymerase to use the new fragment of DNA as a template to do its job. The ligase (which is also present in the reaction!) will then repair the nick that is left once the polymerase has done all it can.
NEB has a great website that explains this whole process very clearly (https://www.neb.com/products/e2611-gibson-assembly-master-mix).
Best,
Dan
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If I get the sense and antisense made with the overhangs I desire; I think (maybe) the homology between the sense and antisense won't interfere with the assembly. 
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Which products? When you combine the backbone and your instert in the 2nd PCR Reaction? This could be possible, if the plasmid is circularisized then it is migrating faster than an linarized plasmid. Essential is that the band of the plasmid backbone is gone, because they are responsible for growing colonies on the plate without insert = false positive colonies. If insertbands are there that makes no effect, the insert has no antibiotic resistance gene.
In the picture I have attached gel slot 2 & 4 show the insert + backbone unamplified whereas gelslot 3 & 5 show a successful joining of the insert + backbone after the 2nd PCR reaction. Here you see that the joined product is also shorter as the theoretic lenght but this is Okay. (Slot one is the FastRuler Middle Range DNA Ladder Marker highest band 5k bp, second 2k pb)
I hope this helped you :-)
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I have used Taq based HiFi polymerase to PCR some fragments with then I assembled in a vector using Gibson Assembly. However, upon sequencing the samples, I see that there are many mutation. I have assembled small (350bp)+ big (nearly 2700bp) together in pBS-SK. The small products were made using normal Taq while the bigger ones using HiFi Taq. Also, important to mention is that the sequencing results have a high background which the sequencing company told me could be artificial sequences because of high GC content. I don't understand what I should do. Since, there are many mutations which is highly unlikely for a HiFi Pol to be going so bad, what should I be doing. Should I assume that these mutations are actually false negatives or something like that.
I would appreciate your help.
Thank you
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Hi Ikram I think if you get a mutant and a non-mutant nucleotide at the same position from different reads I think it is safe to assume that the sequence at that position is non-mutant (and that the "mutant" read is an artefact). If both clones are mutant at the same point then it is safe to assume this is the correct seq (but it may be a polymorphism so you need to check the a.a. seq at these positions).
 It sounds as if you are doing everything correctly so hopefully your new template will give a good seq read (and if the new clones are show mutations at the same positions as your previous clones this indicates these are present in the original template).
Good luck and have a good weekend,
Gary
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Gibson Assembly is a rapid assembly method that provides directional cloning of multiple DNA fragments in a single reaction, without the need for specific restriction sequences. It relies on use of an enzyme mixture consisting of a mesophilic exonuclease, a thermophilic ligase, and a high-fidelity polymerase.The 5' Exonuclease used in this method does not chew the entire DNA.
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It's supposed to get killed by the 50C reaction temperature, after eating some of the DNA.  That's expressed in the word "mesophilic". 
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I would like to assemble a linear fragment together in Gibson and PCR amplify the assembled fragment for cloning. Has anyone tried this? It shouldn't work because the exonuclease will destroy the external priming sites. I was thinking to perform the assembly and later spike the assembly with the external primers. By this time, only the polymerase and ligase will function to fix the ends where the primers bind. Thoughts?
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Hi Rob,
If you want to assemble DNA fragments into one linear fragment, you can achieve this by doing a simple overlapping PCR. Make sure that your two fragments have compatible overhangs, and mix them together with your usual PCR mix, but without any primers. As the strands separate then hybridize, each fragment can serve as a primer for the other as illustrated in the attachment. Calculate the elongation time so that complete elongation can occur. Ideally, make sure you have the same molarity for both fragments in the initial mix.
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I'm using the gibson assembly to assemble 3 fragments (1.5kb 1,5kb and 3kb) into a vector (topo2.1 4kb) to generate a 10Kb plasmid. 2 fragments and the vector are PCR amplified and purified from gel while the 3Kb fragment has been synthesized (gBlock). All fragments contain 20bp overlapping sequences.
I obtained many colonies following transformation but unfortunately I never got properly assemble vectors (digestion analyses showed abnormal digestion patterns with overall shorter or longer size compared to the expected one, as if fragments have not been included or included multiple times).
I tried to change the molar ratios between inserts and vector from 1:1 to 3:1, I also used different bacterial strain mutated in RecA genes, and tried the new HiFi DNA assembly kit from NEB but with no success.
Can anybody give me some advice for successful assembly? Any help is appreciated
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Hi Paulo,
Too weird! I am sure you have already, but have you checked all your fragments to make sure they are correct? Also recheck your overlaps and make sure they are correct. The only time we have had Gibson Assembly fail to give us the correct construct has been an experiment design problem on our part! Most common for beginners (and I am not saying that you are a beginner, just saying that in my experience the first couple of times people use Gibson Assembly, this is where they go wrong) is to have the overlaps reverse complimented to the orientation it should be. Another unexpected issue we had run into was that one of our cloning strategies involves a large binary vector that contained repeats. Having used one of these repeated areas gave one of our grad students a lot of grief.
I hope you figure it out, sorry I can't be more help!
Cheers
Ron
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The Gibson Assembly Mix is assembled on ice for obvious reasons, including not chewing back those ends too far.  However, is there a suggestion to go into a machine pre-warmed to 50 degrees C or should the machine go from 25 to 50 with the samples in it?  This will have something to do with balancing the denaturation rate of the exonuclease versus its activity at 50.
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There seems to be lots of room for deviations.  I pipet mine together at room temperature, mix them, and walk them over to a pre-temperated water bath.  Done scores of them.  They fail due to choosing the wrong sequence overlaps (G/C rich, secondary structures...).  They're very robust against handling details. 
Think about it -- it doesn't really matter if nuclease chews back 25 base pairs or 250.  Pol will fill them back up when it's all said an done, as long as there's a single bp left double-stranded. 
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I have some troubleshooting concerning NEBuilder® HiFi DNA Assembly, and hope someone can give some suggestions. 
I am trying to clone hU6-gRNA spacer-scalfold in a vector around 9kb with this assembly MM. Strange thing is I got many clones (actually not empty vector, nor right clones either ) with very low DNA concentration after miniprep (usually 10g/ul). 
As for assembly protocol, I tried
vector:insert of 1:1, 1:3 and 1:5, with total DNA 0,12pmol,
primer overlap of 20bp.
incubation time 15min and 1 hour. 
Any suggestions?
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Thanks, Ye! Actually I used that vector at that specific site to clone other sequence and it succeeded.