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I am currently generating a genomic rescue construct, and I need some help because it is not working. No-one in my lab has done this before and I don't want to waste all my time re-inventing the wheel. If you have done this before, please give me some advice on how to do this correctly:
I am trying to make a construct consisting of the Drosophila P element vector pCasper4 with an insert of Drosophila genomic DNA that is about 6kB. The source of the insert DNA is a BAC clone. My overall plan is to restrict the BAC DNA, purify the specific DNA that I need by gel extraction, and ligate into pCasper4.
The BAC is 180kB. So far I have:
-Grown up 100mL of bacteria with the BAC clone and isolated the BAC. This was done with phenol-chloroform extraction. I obtained about 20 ug of BAC. The BAC clone is a clean, solid band when run on a gel, without bacterial genomic contamination.
- I cut 1.25 ug of BAC overnight with EcoRI and BamHI (the enzymes i need to generate the desired insert). The resulting digested DNA runs as many distinct bands with a fainter background smear. <Note: I had previously found that 250 ng BAC does not cut at all when digested for 2 hours, with 20 units enzyme each, in a 20 uL reaction. The digest of the 1.25 ug BAC was done O/T in a very dilute reaction (about 200 uL) with 120 units enzyme each.>
- I cut pCasper4 with the restriction enzymes for 2 hours. I gel purified the vector and then cut out and purified a band from the BAC digest between around 5.2-6.7 kB (the band I need is 6kB). I ligated using a 3:1 ratio and plated the entire transformation. I got >200 colonies. I tested 60 colonies, mostly by colony PCR, and some by mini-prep and restriction digest, and none have the correct insert. I think this failed because either too much restriction enzyme was used or too wide of a band was cut out of the BAC digest, resulting in too much background. It is difficult to purify more than 1.25 ug DNA b/c the DNA needs to be relatively dilute to be able to cut.
- For the second try, I cut 1.25 ug of BAC DNA in a 100 uL reaction O/T with 80 units enzyme. I cut out a narrower slice just at 6kB. I ligated and transformed. I got 40 colonies. None had the correct insert by colony PCR. I know the colony PCR is working because the positive control for the colony PCR worked.
If anyone has specifically done this before, please give me your advice. I think some options are:
- cut greater amount of BAC DNA
- use less restriction enzymes (provided the BAC will still cut) maybe there is non-specific cutting b/c too much enzyme is present
- try using different enzymes
I am thinking of not using the BAC at all and using PCR to get the insert.
If I were you, I wouldn't just add more DNA to the reaction. I've come to believe that if you can see it, you can clone it. Are you using the right buffer for your restriction enzymes? For Fermentas enzymes, they suggest 2x tango but the BamHI is reduced to 50-100% activity. What company are the enzymes from? One thing you can do is treat one of your linear fragments with Calf Intestinal Alkalyn Phosphatase (CIP). This will stop re-ligation of your fragment but don't do both fragments (and deactivate the one in the mix) because it will dephosphorylate the required sites needed on your insert for ligation. What was your positive control in your colony PCR?
3 posts • Page 1 of 1
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