Hello,
I am following PCR cloning procedure. I have been attempting to ligate a 700 bp gene into pET28a vector using restriction enzymes NdeI and EcoRI, and transform it into E. coli DH5alpha competent cells. I have prepared the CaCl2 competent cells myself and I am getting decent number of colonies when transforming undigested vector. However, I have not been able to transform the ligated product despite trying repeatedly.
This is the procedure I am following:
Restriction Digestion Reaction: 37 degree C - 30 minutes
Nuclease-Free Water 12 µL
10X Buffer 4 µL
DNA 20 µL
NdeI 2 µL
EcoRI 2 µL
Restriction inactivation: 80 degrees for 20 minutes
Ligation Reaction: 22 degree C - 30 minutes
Vector 10µL
Insert 2.5µL
Buffer 2µL
Ligase 0.2µL
NFW 5.3µL
Ligation inactivation: 70 degrees for 5 mintues
Transformation:
10 microlitres of ligated product in 200 microlitres of comp. cells.
I would appreciate ideas on where the problem might be and how to fix it.
I'm coming into this assuming that you're doing this protocol for the first time and it hasn't been optimised for yourself or your lab. Generally when doing restriction digests I'm quite wary of the timings for "fast digests" or "quick ligation" because of personal tests I've conducted. So I generally stick to a minimum of 2 hours incubation (37C) for restriction digests and 1 hour incubation (room temperature, about 22C) for ligation just to be sure. If I have a lot of time I do both overnight, given that the enzyme doesn't do funny things like show star activity.
Also I didn't see any indication of you purifying your product post-digest, and that can really help with ligating and transforming your desired product efficiently. Just run your product through gel electrophoresis and do a gel band purification so that you get a large majority of the unwanted products out of your samples. If my excised fragment is really small sometimes I just do a cleanup directly using a kit, though running it through a gel gives me more peace of mind.
As mentioned in the above answers, product concentration matters. I usually do a 1:7 plasmid to insert ratio which might be a bit excessive to some people but has worked for me so far. I quantitate using nanodrop but if you don't have access to one you could always roughly put lesser plasmid to insert and try to change the ratios up if necessary.
Hello Arjun
You are not alone with your problem. You simply need to do some controls to pinpoint your issue.
Firstly, you just indicate your volumes of DNA, you must also keep track of your concentrations. A simple way to do so is to monitor everything on agarose gel electrophoresis. Apply samples from plasmid and insert (after restriction) and separate on agarose gels. Test also the restrictions with individual restriction enzymes as well as with both together. Isolate the fragments from the gel and do the ligation. Verify the ligation again on agraose gels with a fraction of your sample. If everything is OK go on with transformation.
Best of luck
You can do the ligation overnight at 16C, the success rate is very high.
You can try out some of these:
- increase digestion time (few hours to overnight)
- run the digests on agarose gel; cut and purify desired bands
- increase the amount of insert for ligation
- try ligation at 16 degrees overnight
- electrotransform or transform using heat shock method
All the best
Some things you should consider:
1. The DNA shouldn't be more than 25% of the restriction enzyme reaction (especially if your DNA is after Clean Up - more of DNA could inhibit digestion)
2. Maybe your insert is toxic for bacteria.
I'm coming into this assuming that you're doing this protocol for the first time and it hasn't been optimised for yourself or your lab. Generally when doing restriction digests I'm quite wary of the timings for "fast digests" or "quick ligation" because of personal tests I've conducted. So I generally stick to a minimum of 2 hours incubation (37C) for restriction digests and 1 hour incubation (room temperature, about 22C) for ligation just to be sure. If I have a lot of time I do both overnight, given that the enzyme doesn't do funny things like show star activity.
Also I didn't see any indication of you purifying your product post-digest, and that can really help with ligating and transforming your desired product efficiently. Just run your product through gel electrophoresis and do a gel band purification so that you get a large majority of the unwanted products out of your samples. If my excised fragment is really small sometimes I just do a cleanup directly using a kit, though running it through a gel gives me more peace of mind.
As mentioned in the above answers, product concentration matters. I usually do a 1:7 plasmid to insert ratio which might be a bit excessive to some people but has worked for me so far. I quantitate using nanodrop but if you don't have access to one you could always roughly put lesser plasmid to insert and try to change the ratios up if necessary.
As suggested by BeNjamin, You must purify your vector as well as Insert before ligation. Digestion is difficult step and more difficult is to determine whether your DNA is being digested or not. Please ensure digestion of your vector and Insert. Checking digestion of insert(PCR Product) is not possible but you can check digestion of Vector. You have also not Mentioned quantity of DNA taken for Digestion and ligation.
First of all, thank you everyone for the inputs.
- I am estimating insert and vector concentrations using a quantifiable DNA ladder. I am getting concentrations of approx. 3 ng/microlit. for vector and approx. 6 ng/microlit. for the insert. Are these concentrations too low?
- As for post-digestion treatment, I have heat-inactivated both REs at 80 C for 20 minutes. I have also cleaned up the digested mixture using a kit. I am also getting size-appropriate band for the vector in agarose gel but I cannot be sure whether both vector and insert are being doubly digested.
- I performed ligation at 16 C overnight as suggested by Marta and Saroj. Today, I performed transformation again (fingers crossed for tomorrow) and also run the ligation on agarose gel. I am getting quite a few bands from the ligation mixture: a faint band for insert, a band at double the insert size (insert concatamer, I am assuming), a band higher than that for digested vector (the sharpest band) and another faint band above that (I am not sure about this one). The DNA ladder I have has a rather low resolution at higher fragment lengths, so I am not sure whether the sharp band I got in ligation mixture was my ligated product or because of band shift due DNA ligase.
- The insert expression is inducible and I have not added the inducer in the selective media. Also insert is not toxic as far as I can tell.
I am sorry I did not respond to most suggestions earlier but all of them have been very helpful. Thank you all. I will let you know if I get the colonies tomorrow. :)
As for the controls Dr. Bülow mentioned, I have set up controls with just comp. cells, comp. cells + undigested vector and comp. cells + digested vector. I got a decent number of colonies using the undigested vector, and no colonies in the other two plates.
Thank you everyone. I got quite a few colonies in my ligation plate today. I haven't screened for recombinants yet but I am hopeful that at least few of them will be.
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