I'm using a dsDNA oligo "Insert" I order from IDT w/sticky ends and 5' phos. I set up a double digestion of my vector as follows:
10x Buffer B (Promega)= 4uL
BSA (10ug/uL) = .4uL
Vector (1.9ug/uL) = 1.05uL (2ug)
---mix by pipetting, then (+):
SpeI (10u/uL) = 1uL
HindIII (10u/ul) = 1uL
Tot. Rxn Vol = 40uL
I incubate this at Room Temp for 3hrs then Gel Purify the band after running on agarose gel. I use the GeneClean Glassmilk beads for this. Everything looks good up to this point. My gel has a single band at the right size for digested vector. I've included SpeI only and HindIII only controls for pMIR vector as well as pBMN and both enzymes are digesting.
For my ligation I'm using T4 DNA Ligase with T4 Buffer. I've been using 100ng of vector and have tried Insert:Vector ratios of 1:1, 2:1, 3:1, 4:1, 10:1, 15:1, 20:1, and 30:1. All of these have been in 20uL Rxn volume and conducted at RT for 3hrs (except 1 where I ran it at 4oC overnight).
I use standard tranformation protocol for XL-1 Blue E.coli: Thaw (+) DNA Rxn directly to E.coli--> leave on ice 20min--> heat shock 42oC 90sec--> ice for 1 min --> (+)300uL LB and shake for 45min @ 37oC then streak on Amp plates and grow up overnight @ 37oC.
I've included positive ctrl of pMIR-REPORT-luc with NO digestion and get tons of colonies. I've done neg control of double digested vector (-) insert and I get zero colonies. I've included a BlpI restriction site in my insert (there's not one in the rest of the vector) for a diagnostic digest. The few times I've gotten colonies (every time its been in 4:1 and/or 10:1 Rxn) the BlpI digestion of a mini-boiling prep does not give me the right band size. Typically I've been getting two bands: one around the right size and one about 3kb smaller.
I have NO idea what's going on. This should NOT be this complicated. ANY thoughts would be appreciated.
Edited by abyrd98, 21 September 2009 - 10:59 AM.