1) Do you use antibiotic selection plate or agar without antibiotic selection for DH5a? Empty DH5a will not grow on antibiotic plates as it do not have antibiotic resistance.

2) Regarding the odd result, have you check if there is any multiple Age1/Xho1 cutting site within the insert? If there is, that would explain why you get fragments of different sizes (smaller than your expected insert).

3) If you use ligation mix to do pcr check, the pcr may amplify from the insert fragments that you put into the ligation mix regardless whether your vector contain the insert or not, so you will get false positive. Normally after ligation, we do colony pcr to confirm if the colonies have the expected insert or not. Using this method can solve the problem stated in no. 2 above.

1. The colonies that have been produced might be from your self ligated vector. You need to choose your selection medium according to your vector of interest.

2. Whether the cloning reaction done at RT before incubating the mixture with competent cells on ice? Usually 15-20 mins at RT before keeping them on ice works well for us.

3. You need to check the antibiotic conc. also, since too high concentration wont give you colonies.

4. As replied by Mr.Viki, you never know if your PCR is positive just because of the presence of unligated DNA fragment.

If you really want to know whether those PCR positive colonies have ligated DNA, try using PCR primers present on the vector flanking your DNA of interest instead of your specific primers. Since you know the bp size of vector primers, you should get a product larger than this if it has ligated DNA, by which you can potentially differentiate.

Thank you Vikki,

1- Our plates have Amp.

2- The fragments used for ligation do not have extra RE sites. We know that because we know their sequences and because they have the expected band sizes on gel.

3- The suggested PCR will have insert and vector specific primers that should read through the cloning site and show that proper ligation happened. Unligated insert should not interfere with that.

Not sure though whether the ligation mix could inhibit the PCR reaction?

Mohamed

Thank you Santhosh kumar Duraisamy .

2- Yes we did 10min and 2h at RT before transforming DH5a on ice, but did not work as mentioned in my original post.

4- We plan to run the PCR with insert and vector specific primers that should read through the cloning site and show that proper ligation happened. Not sure though whether the ligation mix could inhibit the PCR reaction?

There could be several issues for the lack of cloning. The first would be the transformation efficiency, assuming that the "vector only" transformation as your positive control, you should have had higher number of colonies. Have you done the profiling for the "vector only" after the transformation. Although the negative control had no growth, it is still doubtful that your experiment was devoid of contamination as observed during subsequent analysis. How long was it, before you checked the plates after transformation, long period would mean for the growth of pseudo colonies. However, it is still possible that you had contamination during the miniprep culture itself, but the selection pressure would have taken care of that. How much of ligation mixture did you use, since the size of the vector comes to be around 8.2 kb and it is inherently difficult to transform as the size of the vector increases. You could try increasing the concentration of vector and maintain a molar ratio of 1:6 for your next transformation. You could try with 100 ng of vector and 60 ng of insert. Even though the PCR could give you a positive result, it is not a guarantee that you will get the recombinant vector after the transformation.

Hope this is helpful.

It might be possible that your vector was not fully digested and therefore, your ligation is nearly impossible. To improve your digestion efficiency, you might try to do a sequential digestion of the vector, starting with XhoI, purify the product, digest with AgeI, purify again, and then try the ligation. You can try various ratios of insert:vector (molar ratios to be more precise) 5:1, 10:1 or even 20:1. Sometimes only one of then works, sometimes all work, and sometimes nothing works. You can screen your colonies by colony PCR or preparing boiling mini-preps before doing a proper mini. If you get trapped in a situation where after various rounds of ligations your cloning still doesn't work, I would recommend to switch to gibson assembly, which is a cloning strategy independent of ligation.

Given that none of the ligation reactions result in more colonies than your vector only ligation plate, I do not think it is worth your time to further screen the colonies with PCR. If you are concerned about your competent cells or your ligase you can set-up positive control reactions to test these reagents. Sebastian's suggestions to improve digestion efficiency of the vector and and trying more insert:vector (molar) ratios are excellent (I have even done 50:1). Since the 2 enzymes do not cut with 100% efficiency in any buffer this could be an issue with the cutting of both your insert and vector. Do you feel confident about the DNA quantification of the cut insert and vector? Often I use as little as 10ng of vector for a ligation to keep the reaction volume small (10ul) and then use the entire ligation for a 100ul transformation (I usually make my own competent cells, so the transformation efficiency is typically lower than purchased cells). However, given your low yield you may choose to start with 20ng or more vector. Also, using too much ligation reaction can inhibit transformation efficiency (never exceed 10% the volume of the cells, sometimes less is more effective).

Thank you Mitesh, Sebastian and Rebecca for your input. I will try higher ratios and see.

Ligation reaction will not interfere with the PCR. PCR with insert+vector specific primers should show you, whether the ligation was succesfull. You can even run quantitative PCR to see, how efficient it was (in this case, you should run qPCR with vector specific primers only (eg targeted to b-lactamase).

Sometimes, DNA extracted from gel is hard to work with. Partially because of impurities coming from agarose (repeated ethanol precipitation/washing should help), partially because of UV ilumination.

For best results, do not expose your fragments to UV light - separate your DNA to be isolated in EtBr free gel, run a small aliquot of your DNA (would serve as a marker) in a separate lane. After separation, cut the "marker" line, stain with EtBr and identify position of the DNA of interest under UV. Cut the coresponding part of the unstained gel with your DNA of interest.

Restriction-do not overdigest your plasmids, very long incubation times might give worse results.

Dephosphorylate either vector or insert (just one of them), to prevent concatemer formation (if using higher insert:vector ratio, dephosphorylate insert).

Good luck

Thanks Martin for your insight. I will try the PCR using ligation mix following your input.

i agree with Mo Mb

It is possible the transformation was not efficient. Using a high ratio of insert does not necessarily increase the efficiency. You may need to calculate how much insert to use based on the sizes of your insert and vector.

You may try a lower annealing temperature while pcr-screening the colonies. I noticed that it is possible the annealing temperature that worked well when you pcr-purified the insert might not work well during colony screening. I had this experience in the past. For some reasons unknown, primers became more sensitive to high temperature after the first time I used them.