Thanks for posting the extensive information of what you have done so far!

A few potential problem areas for you to check/consider:

1. On your PCR insert, did you leave enough bases before/after the restriction site such that the enzyme is able to digest efficiently? What enzyme site did you use?

2. You should keep the total amount of ligation DNA to under 40ng. Too much DNA overwhelms the cells.

3. Increase the insert:vector ratio. Most people still do the classic 3:1 ratio. That worked poorly in my hands so I typically do 40:1. 50:1, 60:1 & 80:1 ratios and one/two of those always worked (the 40:1 & 50:1 most of the time). You can decrease the amount of vector to 0.5 or 1ng so you don't have to use an excessive amount of insert to achieve the high ratios.

4. Plate ALL of the transformation. Ligations are very inefficient so plating the entire transformation will increase your colony number. After heat shock, add 250ul broth and recover for an hour then plate the entire volume on your plate.

5. Making your own competent cells is always problematic. If you can afford to, buy the high efficiency cells from NEB (don't bother with the subcloning efficiency ones). You can use 25-35ul of the cells to make them last longer and still get good transformation efficiency.

1. That does sound low to me for a cloning strain like DH5a, but I can't really tell you without knowing the volumes involved or the cell density. Transformation with ligated DNA product is always less efficient than a supercoiled plasmid prep so, not a good sign when your competent cells produce way less colonies with a positive control than expected.

2. I prefer electroporation to chemical transformation - but if you absolutely have to use chemical transformation and make your own competent cells then this method: http://cshprotocols.cshlp.org/content/2006/1/pdb.prot3944.long works pretty well.

3. Support for your claim should be "I am wasting time, money and reagents and it's probably overall cheaper to just order new ones." I'm guessing cloning is supposed to be an incidental part of your thesis and not the focus.

There's another potential flaw I can see with your cloning protocol though - excessive gel extraction steps. If you're using UV illumination (instead of one of the blue-light excitable dyes) you're damaging a good portion of your DNA each time you expose it to UV light. Even on low power. Some gel dissolving buffers (depends on the kit) also contain high concentrations of sodium iodide which can get oxidized over time to elemental iodine, which can also damage DNA. If the kit is expired either get a new one ordered, or if that isn't an option try to find if the buffer contains sodium iodide. If it does and the buffer looks yellow - mix in small amounts (like 50 µL to 50 mL buffer) of beta-mercaptoethanol until it turns almost entirely clear again. This will make it a bit stinky but BME reduces the iodine to iodide. Some kits put pH indicators in the buffer which can also make the buffer yellow though, so be aware of that.

Personally how I would approach this if I were you is:

1. Use a PCR purification kit on your PCR product. Do not gel extract it here.

2. Digest with restriction enzymes.

3. Now gel purify your digested product/vector.*

4. Ligate the gel extracted products together.

*First ask if you really need to use gel purification. If you have a single bright band after PCR, you probably don't actually need it so long as the template DNA can't transform into E. coli. You can use phosphatase treatment on the vector to keep it from self ligating. You will still get some background but usually not enough to cause problems.

If you need to do gel purification:

Use a dye like SYBR safe on a blue-light transilluminator, if possible. If you need to use UV, use low power and work as quickly as possible. Use double the amount of wash buffer on all desalting wash steps (the ones after the dissolved gel goes through the column). Let the desalting buffer sit on the column 5-10 minutes before spinning. If the kit only recommends one desalting step, add another. In my hands that gives me very good 260/230 ratios and then I don't have to worry about an additional cleanup step.

Tom Masi

Hello, thanks for the answer. To go through your suggestions:

1. One site only has 2bp exess. I already read about it and I noted it as ruled out that thats causing issues because a) ligation/transformation of just the uncut vector and single cut versions of it also dont work and b) when observing the insert only ligations on an agarose gel I can see fragments above the size I would get if only one enzyme site was successfully cut. By my logic if this were the case only two pieces of insert would be able to stick together. Enzymes are XmaI and MfeI, the XmaI site is added by PCR amplification

2. yes, I actually found that out because I once fogot to dilute my DNA and threw 5ug at them. It was significantly worse than the usual bad results. I'm trying to keep it to 50ng may, but I also noticed that we have unusal bacteria aliquotes of 300ul so I'm not sure if I should scale it up since most trafos start with only 50ul. I've been told the protocol is established and works like it is, however I will consider just setting up a 50ul reaction anyway

3. The highest I'm trying is 1:20, but thanks for letting me know I can just go higher and reduce vector

4. Also a great suggestion I got several times, I will try it next

5. I'm getting close to convincing my supervisors to purchase new bacteria, I need to make sure that they don't die due to problematic handling steps and get wasted. So I'm trying to gather everything I can do better

Thanks for everything!

Alexandra Johnson

Thanks for the suggestions. I'm a little limited on what I can do here, but I think I cut out one more gel extraction using your proposed workflow. I dont have access to things like AP, but I already went from gel extracting vector and insert three times to just once and it significantly improved things.

I will see what I could possibly do with your other suggestions, thank you so much.

It seems clear that the problem is your transformation, most likely the competent cells. Using 10ng of uncut DNA in a typical transformation should give you 10^6 or more colonies (nearly a lawn). If you are getting much less then it is unlikely you will ever recover the product of a ligation. Be sure to confirm that your plates are correct (that someone didn't add the wrong antibiotic or the wrong concentration) but aside from that it is likely the competent cells are not good.

Certainly the easiest solution is to purchase some commercial competent cells, but you might also see if one of your lab mates or another lab has an aliquot of competent cells you can try. Until you get this working well, troubleshooting the cloning steps is not going to be useful.

It seemss that there is an issue with the cells based on your control. I would prepare fresh cells and make sure the competency is high with a control again. When making them keep everything in the freezer (including the containers where you will harvest the cells, wash buffers, tubes where you will store, etc). Also, split the cells in a way that you don't have to thaw multiple times. Do you purify after ligation? If so, have you try EtOH/tRNA ppt? Sometimes you can increase the transformation efficiency with that procedure. It worked for us with a library. We used this method and it worked for us: https://academic.oup.com/nar/article/27/3/910/2902439#.Yo76WV9zsh8.twitter

Hi Louisa, have you fixed this by now? I am facing exactly the same. I have tried everything possible, but there are no colonies after transformation. I have checked my cells, primers, reagents, and everything possible. But this just doesn't seem to work. I had bought the Easy Select kit in February 2021. If you have solved this by now, I would be eager to learn how you have solved it. Thanks!

Hi louisa, I hope you have fixed this by now. I am facing the same problem and I have been stuck here for two months