Hello Abdullah

Are the restriction sites adjacent to each other? I don't know what close they are.

I think, closeness of the restriction sites is the problem you have. If they are too close, one of both enzymes would not cut its restriction site efficiently in the 12.7kb vector, because after cutting of one site, the another site remains without enough extra base pairs to properly digestion. That means, 12.7kb vector was not cut completely

If it is true, you should select another set of restriction enzymes, or another solution is, for example, ligate an insert (a DNA fragment which shares x1 sites in both ends), into x1 site of the 12.7kb vector, then cut the new vector into the adjacent site x2, and finally, cut again into x1 site. This procedure could help you to separate two adjacent restriction sites.

Other suggestions are:

Inactivating CIP, because it would remain active after dephosphorylation step (if you use it).

Using correct restriction enzyme buffers.

Performing sequential digestion reactions.

I hope all this helps to solve your problem

There are many potential causes that may be leading to unsuccessful cloning with the strategy you've outlined here. I would focus on the following as the most likely culprits:

1) your double digests of the vector are not working. This could either be due to the fact that your enzymes are not compatible in the same buffer, or that the cut sites for these enzymes are too close together in your plant vector (please specify the bp distance between these sites in the MCS). In addition, certain enzymes, most notably SalI and StuI, show extremely poor digestion efficiency against supercoiled DNA templates. Furthermore, StuI and ApaI are blocked by overlapping dcm methylation, which could be a potential issue for efficient digestion of your plant vector. ApaI also works best at 25C, and is inhibited by salt >50mM, so your need to be very careful that all your digestion conditions are appropriate.

Since your DNA ends are not compatible, but your no insert, no CIP ligation control gives too many colonies, you can definitively conclude that you are not getting efficient double digestion of your vector.

2) Heat inactivation of T4 ligase and purification of your ligation is not standard operating procedure for cloning, and you risk losing too much material in this step. I would recommend removing these steps from your protocol.

3) Ligation work best with enzymes that leave sticky ends that have a 5' overhang. StuI is a blunt end cutter, while ApaI leaves a 3' overhang. The choice of these enzymes will reduce cloning efficiency. In addition, excessive UV treatment or contaminating salts/agarose in your gel isolation can strongly inhibit ligations. Make sure you use minimum UV exposure to isolate your DNA and check the quality of your gel-isolated DNA on a spectrophotometer (preferably a Nanodrop). Lastly, remember that less is more in cloning (don't put more than 100ng TOTAL (vector + insert) DNA in your ligation reaction). Too much DNA can decrease transformation efficiency and promote excessive intermolecular vector-to-vector dead-end ligations.

Hello Javier Montalvo and Daneil Cohen

I am really appreciate that.

Daneil Cohen

The clear map for the reconstructed pCAMBIA vector (plant vector) is not available to me however the restriction map shows that the MCS is about 108 bp have 17 restriction enzymes and ApaI SalI are so close, may be overlapped, . So, I had another option is to cut pCAMBIA vector with ApaI only (linearized) and cut pUC57-X1 vector with ApaI only (it give 2 bands one shows the correct size of my insert), so that I could ligate the lower band into single cut vector, after treating the linear vector with CIP, but I got no colonies at the end. Could you consider that.

Overdigestion with CIP or unsuccessful removal of CIP will prevent successful ligations. Make sure you use no more than 0.5 unit of CIP per microgram of DNA and digest for no more than 1hr. If ApaI and SalI overlap in the pCAMBIA vector, you cannot digest with both enzymes. For the ApaI only strategy, do you get no colonies at all or no recombinant colonies. The distinction matters for troubleshooting.

How much DNA are you using in your ligations?

For ApaI cutting only, after ligation and transformation , I got about 8 colonies, 7 of them contain lower plasmids that often come out (2-3 Kb in length) and one show pCAMBIA as empty vector with no insert after cutting with ApaI only.

For DNA I used different ratio of vector:insert, the last times I put 1.5 microliters Vector (about 130 ng), 3.5 microliters Insert (about 60 ng), this is 1:3 ratio.

I did other reaction using 5 microliters vector to 3 microliters in 20 microliters reaction.

Actually i have trouble understanding the different between DNA concentration and molar ratio and calculating how much DNA should be added in ligation reaction. I used to measure the concentration based on how it is appear on the gel, then compare band strength of vector and insert and take double amount of vector to insert.

After that, I use a software : http://django.gibthon.org/tools/ligcalc/. So, i don't have a clear idea about.

Does the 2-3kb size match the parental pUC57-GOI vector? If not, it is a bad sign that you are getting contamination by a random plasmid.

Estimating DNA amounts by ethidium bromide staining intensity on an agarose gel is harder than reading directly on a spectrophotometer. Do you have access to a Nanodrop? If so, you should be reading DNA concentration by A260.

Once you know the concentration of your vector of insert and vector, you should proceed as follows. Use 25ng of pCAMBIA vector in a 20uL ligation. The amount of insert should be 25ng * size of your insert in bp / size of the pCAMBIA vector in bp * the molar ratio.

So for example, if you want a 1:3 molar (vector:insert) ratio with a 1 kb insert, you need 25ng of pCAMBIA and 25*(1000/12700)*3 = 5.9 ng of insert. You'll need to convert this to a volume based on the concentration of insert (for example, if your insert is 20ng/uL, you need 5.9/20, or about 0.3 uL volume). Make sense?

Based on a 2kb insert size, 130ng of 12.7kb vector, and 60ng of insert does correspond to a 1:3 molar ratio. However, your total amount of DNA in the ligation should be

I agree with Daniel. The UV damage might really be a problem with large fragments. The larger the fragment, the smaller percentage of the DNA which doesnt get crosslinked even by few seconds exposure to UV. Either cut your slices over an additional glass plate which will block the hard UV irradiation, or skip the gel purification completely. I have been cloning a lot into plant virus veotors of about the same size lately. I do a double digest of the vector, dephosphorylate it with shrimp alkaline phosphatase and only check a small aliquot on a gel to see if everything seems OK. Then I deactivate the enzymes and optionally purify the fragment from solution and ligate it with the insert.

You also mention isolation of small 2-3 kb plasmids from your colonies. This might be a problem with plasmid stability. I have had this problem with pGreen based vectors. If this is the case you might want to try different strains of E.coli (Top10 worked better than DH5a in my case), lower the temperature from 37 to 28 and lower the antibiotic selection - i went from 50 µg/ml Kanamycin to 25. What also worked for me was to transform the ligation directly to electrocompetent Agro, where the copy number is much lower and the plasmid is stable.

Daneil Cohen

This is an update for the cloning work

No colonies were observed on all plates.

I cut the pCAMBIA vector with BstBI/StuI (about 12 bases separate the 2 restriction sites), turn to be linear vector, and cut pUC57-FAR with the same enzymes to get my insert out. After digestion i use 0.2 ul (5 units) of CIP to dephoshorylate linear vector, then inactivate CIP by incubating at 70 C for 10 min. I ran a gel to check the digestion and cut gel slices and purify using ZymoResearch gel purification kit. I ran a gel to check the concentration of purified fragments on the agarose gel. I measure the concentration using Nanodrop 2000, it was, the linear vector A260 = 48 ng/ul, A260/280 = 1.71, A230/260=0.32. The insert A260 = 22ng/ul, A260/280 = 1.85, A230/260=0.03.

for ligation I used 1.5 ul vector : 1.5 ul in 20 ul, and 2.5 ul vector : 2.5 ul insert in 20 ul reactions. Incubate tubes at 16 C for 16 H.

For transformation: I used 8 ul of ligation mixture into 50 ul ECDh5alpha.

The Time Constant was too low, 1.3 and 2.0, but i didn't notice a sparking.

I used 950 ul LB to recover transformed cells at 37 C for 1 h and then spread on Agar plates contain Kanamycin 40 ug/ul. I spread 100 ul and 300 ul on plates. Incubate the plates overnight at 37 C.

No colonies were observed on all plates.

Try without the CIP treatment. I usually use heat shock transformations, so don't know off-hand whether your electroporation conditions are appropriate. Often using less of the transformation mix, and pre-diluting into water helps. Also, you should probably use a ligation buffer that includes PEG to promote ligation efficiency at low DNA concentrations. Just to give you a feel for DNA amounts, I typically set up 20uL ligation reactions with 25ng of vector, dilute 1:5 in water, and use 2uL for transformation (equivalent to 0.5ng of vector). Too much buffer and/or DNA from the ligation mix can inhibit transformation efficiency.

Hello Hesham Abdullah

After DNA purification from gel, you had a low A230/260 ratio in the DNA samples (0.32 for plasmid, and 0.03 for insert); I think that DNA samples are contaminated by salts or solvents. This kind of contamination decreases the A230/260 ratio and could decrease ligation efficiency. Re-purifying the DNA samples with ethanol precipitation could work.

Daniel Cohen

Here is un update.

Exp.1 :

I've tried to open the pCAMBIA vector with ApaI and use ApaI to get my insert intact from pUC57 vector, then I did two treatments for cut pCAMBIA vector with and without CIP (CIP treatment: 0.2 ul or 5 U to the digestion mix and incubate for 1 h at 37 C, inactivate by adding 0.5 ul 0.5 M EDTA and run a gel ). I did two ligations, in 1st one i added 3 ul (30 ng/ul) of cut, no CIPed purified vector + 2 ul (23.3 ng/ul) purified insert in 20 ul reaction.

2nd

I added 2 ul (49.9 ng/ul) of cut, CIPed and purified vector + 1 ul (85.5 ng/ul) purified insert in 20 ul reaction.

Incubate the reactions for 16 h at 16 C. I use all 20 ul transformed by heat shock using 50 ul Dh5 alpha in Ice for 30 min, at 42 C for 90 seconds, then in ice for 5 min,

Results, 1 or 2 colonies were showed per plate, however it didn't grow in overnight suspension culture in the presence of 40 ugm/ul Kanamycin.

Exp.2 : I've tried to open the pCAMBIA vector with SalI only, CIPed and gel purified and use SalI to get my insert intact from pUC57 vector, then. I did two ligations, in

1st one

I added 2 ul (80ng/ul) of vector + 2 ul (60 ng/ul) purified insert in 20 ul reaction.

2nd

I added 4 ul (80 ng/ul) vector + 4 ul (60 ng/ul) purified insert in 20 ul reaction.

Incubate the reactions for 16 h at 16 C. I use all 10 ul transformed by heat shock using 50 ul GC10 strain (suitable for large plasmids) in Ice for 30 min, at 42 C for 90 seconds, then in ice for 5 min. I transformed 10 ul into 50 ul electrocompetent Dh5 alpha (Time constant was 3.8).

Results, no colonies were showed on heat shock plates, however one colony showed on Electroporation plates. I picked it up and setup overnight suspension culture in the presence of 40 ugm/ul Kanamycin. then I did plasmid miniprep then run a gel. It gave a lower size plamid didn't turn to be linearized when cut with SalI only comparing to linear positive control.

I need someone to explain what is going on and please find the attachment of vector maps I am using

Hesham- did you compare cut vs. uncut vector on an agarose gel to check the efficiency of your ApaI and SalI digestions?

Yes, Idid. For pCAMBIA vectors cut with either ApaI or SalI it showed to be linear (shifted up) while the pUC57-WSD1 and pUC57-WS cut with either ApaI or SalI, respectively, gave two bands similar in length. I purify the lower bands that contain my inserts. The uncut pUC57 showed at 5000bp, while the cut showed one band at 2500 bp and the other band at 2100 bp

It sounds as though either your ligase buffer or your ligase enzyme is dead. You can try to supplement with fresh ATP, but probably the best option would be to get entirely new ligation reagents (both buffer and enzyme). If possible, run a positive control ligation in parallel to confirm your ligase is active.