This may be a simple question but what selection marker are you using for your plates? It seems like both the positive and negative plate has a lot of colonies.

Could you tell us more about which plasmid you are using, what is your insert, do you phosphorylate and anneal the insert, how do you do the transformation?

Hi there,

You should have 2 control plates: one with the vector alone without ligase (to evaluate the presence of intact plasmid in the ligation mix) and the other with the vector alone with ligase (to evaluate both the presence of intact plasmid and the contribution of self ligation). Your control is the latter which makes no distinction between intact plasmid and self ligation... Therefore you still have no clue on where the problem lies (digest and/or ligation)... BTW how many clones did you test? You PCR screen seems to be all or nothing (the set of primers is insert specific) so the absence of band makes no distinction between failed PCR and absence of insert... The best would be to use a set of primers specific of the vector to observe a band shift of the PCR product when cloning is successful.

There's no polite way to say this -- Cloning with restriction enzymes and ligases is obsolete. For those of us who just want a plasmid to be made, this is the worst, least reliable, and most expensive way to go about it.

The best, fastest and cheapest method is to have the plasmid made in its final form by a synthesis company, gRNA, cas9, promoters and all, sequence guaranteed. If you tally up the cost of all your ligation kits, enzymes, and let alone your time, total synthesis is simply unbeatable. Just don't make a mistake in specifying what the sequence should be.

Optionally, if you want to learn how to clone (or your PI insists that you do), then do your future self a favor and learn a modern method that actually works most of the time, like Gibson's Assembly.

You used single restriction enzyme for linear the vector. In this condition their is a chances of self ligation of of vector to produce false positive colonies.

Try double digestion of vector and insert with 2 different RE enzyme that eliminate the self ligation.

1) check you antibiotic stock concentration and prepare fresh working stock if possible.

2) Make sure the the your competent cells are not growing in media with antibiotic to check for any contamination.

3) To check contamination in competent cell used 2 culture tube one with only media+antibiotic, media + CC and media+antibiotic+CC.

4) For transformation used a extra control without ligase.

5) While plating make some control

a) CC without any ligation mixture.

b) CC with ligation mixture(no ligase)

c) CC with proper ligation mixture.

I hope this will help you or you will defiantly find a solution for you problem.

Adam Desrochers the plasmid we are working with bRA90 that contains an Amp selection marker. It is fairly big in size 11.8 kb. It also contains a LEU2 selection marker.

My insert is a small segment of the RPT6. The insert is 20 bp in size. So it is fairly small in size. The insert was duplexed using 100µM of each oligo and incubate in PCR at 100 degree C for 5 minutes and cooled down at RT.

I tried using both XL-1 and Beta-10 competent cells for transformation with both giving the same results. The ligation mix was set at RT for 1 hour before transferring 10µl to 100µl of competent cells. I then incubated for 15 minutes on ice before heat shocking cells at 42 degree C for 90 seconds. Cells were placed back on ice for 5 minutes before recovery. I recovered the cells using SOC at 37 degree C for 45 minutes. I plated cells on LB + Amp plate that had been pre-warmed for 30 minutes.

Dominique Liger The vector was treated with rSAP so I assumed that self-ligation would not occur (that's where I am wrong where I should not have assumed). I will try again to see if it self-ligation is occurring. For diagnostic purposes, I would check 12-20 colonies from the plate and pick a colony from the control plate to use as a negative control. The primers that I use to check is my gRNA forward primer and Amp reverse primer.

John Schloendorn I understand where you are coming from. I am still working on my cloning skills. I think it would be nice to feel comfortable with having this technique under my belt. The plasmid we are using contains the Cas9 and promoter already, we just needed to insert our gRNA. The protocol I am using to clone has been seen successful in other labs and it is still being used. Also, we have the reagents in our lab to conduct this experiment.

Des'Ree D Groover I have done a very similar experiment using px459 plasmid. Basically the same thing to load in a sgRNA. I would recommend you try the follwing things:

1. Double check that you are using an appropriate antibiotic concentration and that the ampicilin is fresh. It degrades relatively fast. 50 µg/mL should be sufficient if fresh for plates. When making the plates make sure not to add the ampicillin while it is too hot, this could denature it.

2. Try digesting your plasmid for a longer time to help improve yield and eliminate any non digested fragments. If you are only cutting out a 20bp piece differentiating cut from uncut plasmid is rather difficult on a gel when isolating it. Perhaps a longer digestion willl reduce the number of uncut plasmid in the mix.

It really seems to me there is a problem with the digestion of the plasmid or selection of the plates. If the digestion is incomplete it makes sense that you see many colonies on the negative, likely having received a blank undigested plasmid. I had a similar difficulty before using a similar approach and it was resolved by increasing digestion using BbsI from the recommened 15 minutes from the manufacturer protocol to 2 hours.