More PCR cloning problems! - (Dec/06/2006 )
I've been reading this forum, searching for a solution to my cloning woes, but to no avail! For two months I've been trying to clone a 300bp PCR product into the vector pET17b. I'm using primers that have incorporated restriction sites (NdeI and BamHI), cutting both insert and vector with the enzymes and trying to ligate, but I usually get no colonies. (The few colonies I have gotten have no insert). Here's my basic protocol:
-make lots of PCR product, purify with Qiagen spin kit
-digest insert and vector in 50ul for 2-3 hours with NdeI and BamHI in Bam buffer (I've tried co-digestion as well as sequential)
-i have tried to CIP the vector and also to skip this step
-run digested vector and insert out on a gel, cut out, and use sigma kit to purify (i can see a difference between my cut and uncut vector, but not really for the insert)
-spec. vector and insert, and use T4 ligase from NEB to ligate
for the ligation, I have tried numerous (seems like endless) combinations of insert:vector ratios (1:1, 3:1, 6:1, 50:1, 100:1). I've also tried different incubation times from an hour to overnight at 16 degrees. I thought that the buffer may have degraded from multiple freeze/thaw, so I've tried to supplement with ATP (1mM) as well as using a fresh NEB buffer 4 + ATP.
- transform into chemically competent DH5 alpha cells, usually 5ul ligation into 50ul cells (is this too much?)
- as i said before, i get maybe one or two colonies, but when i check they just look funny and are very high molecular weight, no insert : (
as far as transformation controls go, i do a positive control with uncut vector = lots of colonies, and i do a control with cut, unligated vector = no colonies.
i've checked the ligase, and it seems to work when i cut the vector with an enzyme that cuts three times i can follow the same preparation steps and religate the pieces....although i had to add a final phenol:chloroform purification to get the ligation to work.....if I use a kit to purify after gel extraction, is this step necessary? i have eluted in water and the elution buffer without seeing a major difference.
as far as the primer design goes, they were given to me by the previous PhD student who couldn't get the ligations to work, but a postdoc has used them and they supposedly work.
I'm so frustrated, and I can't do anything until I get these clones to work and make some protein! Please, I would appreciate any suggestions, or if I'm doing something silly wrong, please say so! Thanks in advance for any replies.
Cheers!
I would suggest you omit the gel purification of your cut PCR product. If you have a clean band, you should not need to do this step, and you may be trashing your DNA by exposing it to short wavelength UV during this process. You can test the ligation efficiency of your cut PCR product by ligating it alone, heat killing the ligase, and then running the ligated product on a gel. You should see high MW bands if things are working well. You can test cutting/religation of each enzyme independently and observe a double length band. The relative proportion of single and double length fragments will tell you the efficiency of the cutting and religation.
Hmm.... adding to phage434's suggestion, a look see at the primer design might be needed. The primers already have a bad history, so could we have a look at the primers.
Thanks for the suggestion about skipping the gel extraction step. I'll try that and the cutting/religasing of my PCR product.
My primers are:
fwd: 5-CTGC CAT ATG TCT GCC TCC CCC AAA CAG CGG CGC-3
rev: 5-CCT TTT GGA TCC TCA GCT CCC TCT CCC AGT TAC-3
There is an Nde I site in the fwd and BamHI in the rev.
Thanks!
Just an update...
I tried phage434's idea of digesting my insert with each enzyme separately, and then ligating. Basically, I digested, cleaned up with a PCR kit, and ran it out on a gel. Each digestion looked fine, and I did see high MW product, but it was much greater than 2X my insert. How is this possible? If it's only cutting on one end, then only one end should be free to religate, right? So 2X is the max. size I should see....am I wrong? Both ligations appear to be the same MW. Hmmm....comments?
Are you using quick ligase or normal T4 ligase? The quick ligase buffer contains PEG which should not be heat killed, and will do strange things to your DNA and gel image if you heat it. Unless you heat kill the ligase (in normal buffer) the ligase remains attached to the DNA and gives inaccurate length readings for the ligated product. How are you doing the ligations (temperature, time, buffers, enzyme?).
according to NEB's technical guide, the NdeI site on the forward primer :-
fwd: 5-CTGC CAT ATG TCT GCC TCC CCC AAA CAG CGG CGC-3
has a cutting efficiency of zero ( 0% ). Basically PCR products made with this primer will not cut with NdeI. Only 4bp of guard sequence (CTGC) was added to the 5'end of the fwd primer. NdeI requires a minimum of 7bp on either end of the restriction site to work.
My suggestion is to concatemerise your PCR inserts, (PNK treatment, then blunt end ligation) followed by through digestion with NdeI and BamHI.... which will give you your digested inserts... (though you must be warry of double inserts, when you screen for your insertions.)
Thanks for the suggestions. I was wary of that Nde primer, but I thought that too few flanking basepairs just reduced the cutting efficiency. I will definitely give the concatemer approach a try.
Yesterday I used Quick Ligase at room temperature for 5 min. according to the manufacturer's instructions. I did not heat kill the ligase.
Hopefully it's just a digestion problem and this will be the solution!
Now that I think of it, I've never concatemerised before. Any suggestions for a protocol?!
set up a quick ligation and add 0.5ul PNK. (If your reaction volume is more then 40ul add 1ul PNK) Incubate for about 2hrs at room temperature ~25 Celcius.
After that, you can check if the ligation has worked, by taking an aliquote and running said aliquote on a gel. You can heat kill it first if you want, before running the gel.