Hi, you should get rid of colonies in vector only plates. Not sure about your protocol, but it may be 1) incomplete cleavage of vector (improve this or even gel-purify linearized vector) 2) religation of empty vector (provided the ends are compatible) - dephosphorylate vector before ligation 3) cross contamination. This should help. However, to speed it up, I d recommend to go to different cloning method (such as Gibson assembly), which is far more efficient and quick. If you prefer restriction based cloning, consider PCR based linearization of the vector (nowadays the polymerases can easily do this). The original circular vector can be easily removed before transformation by DpnI treatment. Good luck

Have you tried assesing if your insert ligated succesfully by doing colony PCR, restriction digest from miniprep?. Judging based on the number of colonies might not be accurate. What you need is to confirm it if your insert is indeed ligated.

Hi Ridzky, I didn't go for Colony PCR and or RE for checking the insert ligation considering the no. of colonies are same. Do you suggest that I go ahead and check for the insert in the colonies on my 1:3 and 1:5 plate?

You could try treating your ligation product with T5 exonuclease before you transform. That has worked for me in the past.

Hi Sarmishta. Yes, you can try on the colonies found in both plates. You can try randomly pick 5-10 colonies on each plate and check the presence of insert by colony PCR. I just forgot to ask, during your cloning, did you remove the phosphage group of the vector by alkaline phosphatase? this usually works to prevent vector religation and increases the chances of having insert in the vector. But, please do not treat your insert with the phosphatase, otherwise you will not be able to ligate it. Which enzyme were you using for the ligation? I used T4 ligase from NEB.

I did not treat the vector with AP. I'll try that going forward.

Regarding the ligase enzyme, I used T4 ligase from Thermo (Rapid ligation kit) and kept overnight at 4deg

Look, if you have a colony that has your insert, then you are done.

No need to trouble-shoot the question of "why are the efficiencies different". It's a waste of your time and resources to do any follow up.

Done is done. Move on with your project.

Hi Katie, I agree with you. I'll be checking for the insert. Let's hope to get a positive colony.

Hi Ridzky Yuda I checked for insert in 10 colonies and I couldn't observe the insert band in any. I am planning to set up the experiment again. Can you suggest some dos and don'ts?

Hi Martin Lenicek

I am attaching my vector and insert digestion gel image and I suppose the digestion is complete. Can you please check?

Hi Sarmishta Majumdar , not sure, what is vector and what is insert. Also, which restrictases did you use (single or multiple)?

The vector is my own design, its a mammalian expression vector and the insert is a gene expressing a protein. Two restriction enzymes are used AgeI+XmaI. The vector has a MCS site flanking these RE site for ligating the insert.

Lane 1: ladder

Lane 2: Uncut vector

Lane 3: empty

lane 4: Double digested expression vector (green labelled box)

lane 5: Uncut cloning vector containing insert

lane 6: Double digested cloning vector with insert release (0.7 KB green labelled box)

The double digested vector (lane 4) and 0.7KB insert (lane 6) is purified and used for ligation

Sarmishta Majumdar ok I see. The trouble is, that your double digested vector has very similar migration pattern as the uncut one. As a result even gel-purified vector will not be 100% linear (digestion is never complete). Presence of even trace amounts of undigested vector will result in plenty of colonies containing empty vector (and you will need to screen many of them). If you can not separate uncut and linearized vector, try to minimize the presence of that uncut (perform digestion with tiny amount of the plasmid). Make sure, that your restriction works: are the restriction sites separated enough to allow double digest? Enzymes do not cleave efficiently clost to the end of DNA. If only single digest occurs, your insert would not fit in, but religation of empty vector is going to be highly efficient.

Try to analyze your ligation mixture before transformation - this may save a lot of time - you can run PCR on the ligation mixture: one with primers flanking the MCS (positive PCR - short product indicates presence of unwanted circular vector), second with one vector specific and the second insert-specific primer (positive PCR shows, that you have desired construct in your mixture and that it makes sense to do transformation). If you can run quantittive PCR, you can even estimate the proportion of empty vector vs correct construct.

If empty vector is still present in your ligation mixture, check, if there is a possibility how to eliminate it - there may be unique restriction site in MCS, that is removed during double digest (ie is located between AgeI and XmaI). In this case, cleave your ligation mixture before transformation (you should get way less colonies, but the proportion of the correct ones should be markedly higher).

Still, I´d consider PCR based amplification of the vector - it is fast, easy, you are sure, it is linear and after ligation, you easily remove circular vector (that is methylated) by DpnI treatment.

Martin Lenicek Do you think there might be a problem with the plasmid isolation considering the uncut and cut plasmids have similar patterns on the gel?

The RE sites are well separated (around 250bp stuffer between them)

Sarmishta Majumdar This seems not to be isolation trouble. It is rather failure of separation of doubledigested vs uncut (or singlecut) vector. Both uncut and single-cleaved plasmid will dramatically increase the number of empty clones. You need to solve this otherwise you´d need to screen thousands of colonies.

Sarmishta Majumdar 250 bp separation is good enough for efficiend double cleavage - good news. Still it seems difficult to gel-separate linearized vector from that uncut(your native vector just moves in a similar fashion). Is there a unique RE site in that 250 bp stuffer? If so, use this enzyme to digest your ligation mixture (after ligation and ligase inactivation) to get rid of empty vector before transformation.

The primers flanking MCS that you suggested, did you mean to design primers (one end having AgeI another one having XmaI) for the section of vector starting with AgeI ending with XmaI?

I have a 250bp spacer between these 2 Re sites. Do you mean the each of the forward and reverse primer will be flanking both RE sites? And the in the in between sequence will be of the spacer?

Not sure which primers do you mean. Check the pictures attached. First one shows primers for PCR amplification of your vector. After amplification you just digest it with AgeI and XmaI, ligate and then treat with DpnI and transform.

The second picture shows primers for colony (or ligation mixture) screening for correct clones. Primer combination 1+2 yields short product in empty vector and no product (or very large product) in correct clone. Combination 2+3 shold give you the expected PCR product in correct clone only. Design the primers so that yo get small PCR product (lets say 200 -400bp), as large products may not be obtained during colony PCR (where you use just a suspension of your colony as a template, instead of isolating plasmids).

Hi Martin Lenicek Could you please explain the vector amplification part? Just to double check, you are suggesting to design primers starting from the MCS region towards the RE site and amplify the vector? Post that DpnI digestion?

Also, as I mentioned earlier that I have a 250bp spacer between AgeI and XmaI. While large scale restriction digestion, I observe a faint 250KB band released at the bottom of the well. Does this suggest that my vector is digested properly?

Regarding amplification: yes, i suggest to design primers in the MC (facing out) to PCR amplify virtually the whole vector (except from a small part inside MCS). You will get linear vector, that contains your RE sites, that can be used for cloning (provided you prefer to stick to RE based cloning over preferred Gibson assembly). DpnI cleaves methylated DNA (ie isolated from bacteria) but not that PCR amplified. This helps you to eliminate original empty plasmid.

Regarding digestion: presence of 250 bp band suggests, that at least part of your plasmid has been cleaved. But this is never 100% effective and at least small fraction of uncleaved vector will markedly affect your result. Most of your clones will be empty.