I'm baffled.
On numerous occasions now, I have failed to gain cells on an LB amp plate from transformed TOP10 E. coli cells.
I have attempted various strategies to figure out what the problem is:
My insert gene is small (~200bp) and I normally get low yield, thus I have tried scaling up by restriction digest and I have attempted a PCR of my insert prior to restriction digest to ensure there is plenty of DNA. This works, I get a much brighter band showing up on an agarose gel, at the correct size for my insert too.
I have also ran the the products of single digests, double digest and undigested insert on a gel to show that both enzymes are working effectively - and they appear to be, whereby in the double digest I get the product I am looking for to purify and ligate. I have also done this for the Ppinkalpha-HC vector that I am ligating into, to show that this is also cutting with the enzymes.
I have tried eluting the products from the gel purification into sigma water and elution buffer.
I have tried various ligation ratios.
I carry out my ligations at 4C overnight, but I've also tried 15 min room temp.
Most puzzling of all is that I have attempted to carry out this work alongside a colleague who is doing the same work with another gene (theirs is ~400bp). I did this to ensure I wasn't making any silly mistake or anything. Thus we followed the exact same protocol (even use the same RD enzymes), at exactly the same time, alongside each other.
Low and behold, they gained some colonies on their insert+vector plate, whereas I gained none. We both also ran a control with just the vector in the ligation mix (no insert gene) and transformed this. I gained now colonies, whereas my colleague gained a lot of colonies, suggesting the vector had self ligated and got into the cells, providing ampicillin resistance and allowing colonies to form on the LB amp plates. Although colonies on her negative plate are not desirable, this showed that the ligation is working. My negative ligation sample is the exact same mix, yet I gained no colonies. Given that my colleague did gain colonies, there's liklihood that the same self-ligation of the vector occurred in my negative control, which makes me think my ligations should be working. Thus perhaps it is the transformation that is not working?
But why, given that I carried this out alongside a colleague, following the exact same protocol.
Cells were not killed by the sterilised spreader, it was allowed to cool. Cells are new. The cells are bought in from life technologies already chemically competent.
Some other details:
I heat-shock cells for 30 seconds at 42C.
Ligation mix material works (as shown by my colleagues positive results).
The gene I am working on is very similar to two other genes I have previously been successful with, the only difference being 3 amino acids in the expressed protein sequence. That's the other thing that puzzles me, I did not run into this problem with two other very similar insert genes, which had the same restriction digest sites, etc.
If anyone has any suggestions, I'd really appreciate any help and advice!
After heat-shocking, 250ul of SOC media gets added to the cells and the cells then incubated at 37C for 1 hour. After that, they get spread on the plate (I've tried using the full volume of cells and using half the volume of cells, same results for both - no colonies forming). Another bit of information: when I've added the ligation mix to the thawed cells, I leave them sitting on ice 15 minutes prior to heat-shock (and I've also tried leaving on ice for 30 minutes).
We do use the same competent cells, yes, from the same batch ordered from Life Technologies.
One thing I was thinking of trying was adding mercaptoethanol to the cells prior to heat-shock. I.e. add mercaptoethanol before I add the ligation mix to the cells and leave them to sit on ice for 15 minutes, before heat shock. My reasoning being that it should make the cells more porous/leaky and help facilitate getting the ligated insert/vector into the cells when heat-shocked.
That's provided the transformation is the problem. I was thinking about doing a PCR of the ligation mix using primers that bind to vector sites flanking the insert region. This should provide a band on a gel larger than that of vector only, provided the insert has ligated.
I use a 6:1 ratio of insert to vector. With a final 30ng of vector in the ligation mix. The ligation reaction is carried out in a 10ul volume and all of this goes into a 50ul volume of chemically competent TOP10 cells.
As a side note, I've also tried other ratios, including 3:1, as well as simply much higher amounts of insert DNA in an attempt to improve the chances of insert ligating with vector.
I believe the problem is you are transforming too much volume (10ul) into your cells thus changing the competency of them. As a rule of thumb you should not transform more than 5ul of a ligation into 50ul of cells. I typically transform 1ul of a 10ul ligation into 50ul of cells and have no issues. So I suggest you try transforming less volume and see what happens. If you want to transform that much volume then you should increase the volume of cells you are using. Also, transforming too much DNA can result in no/low numbers of colonies. Less than 40ng total (vector + insert) should be transformed.
David
1. Are you inactivating T4 before transforming? (That always worked better for me).
2. Are you using the necessary volume of T4? Usually they recomend 5U/100ng DNA. If you are using 210ng total DNA (6:1 ratio with 30ng vector) you should need more ligase.
3. I prefer to use molar ratio when cloning small fragments, specially since your vector is huge (40x your fragment). I always calculate 5:1 molar ratio considering pmol ends but keeping a maximun of 100ng total DNA. I have cloned smaller fragment with this method and I like it better because you don't waste DNA.
4. How are you quantificating your DNA? I'm old school, I always had better results by quantificating my DNA with EtBR gel densitometry.
David, I agree with all said here, using too much for transformation, 1-2ul shall be enough and no more than 5ul, use molar ratios, you are using way too much insert, at a 1:1 amount you are about 20:1 molecular ratio ( counting 4 Kb vector vs 200bp insert ) and yes adding a 65C at the end of ligation does help. Best of luck, doing those changes shall work. Mario
Hi all,
First of all, thanks for all your advice and help, I appreciate that.
Just to clear up, as I realise now how unclear I was above. I do use a 6:1 molar ratio. The final amount of DNA in the sample tends to end up around the 36ng mark based on the size of the Ppinkalpha-HC vector and the size of my insert. Apologies for making it sound like a 6:1 ratio based on the amount of DNA in ng.Also, in response to a previous question, I use 5U of DNA ligase in the ligation reaction - thus enough for up to 100ng of DNA as previously stated.
Good news! I went ahead and used 1ul and 5ul of my 10ul ligation mix (carrying out a 6:1 ligation reaction as normal). I have colonies on both plates, a few more on the 5ul plate than the 1ul plate. Since I did nothing else differently, it appears we have cleared up the issue - that I was using too much of the ligation mix in previous transformations, rendering the cells incompetent. Unsure why my previous transformations of other genes have worked with the full 10ul ligation volume (as there should be the same amount of impurities (salt, etc) to my recent attempts at transforming cells with 10ul of the ligation mix and the total DNA in the 10ul ligation mix is still only ~ 36ng.
Thanks again for all your useful help.
David
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