I am trying to label a 2.5kb DNA fragment (~50% GC content) with the DIG High Prime DNA Labeling and Detection Starter Kit II (Roche, cat. no. 11585614910), but my labeling efficiency is 10-20 fold less than the expected yield as shown in the kit manual. I have tried hybridizing to my target DNA and have achieved signal, but it is extremely low. Has anyone had similar issues? Here is the experimental setup:
1) PCR amplify probe, run agarose gel/excise band/purify with Promega Wizard SV kit, ethanol precipitate, resuspend pellet in nuclease free H2O.
2) Follow labeling procedure as outlined in kit manual: Boil DNA (1ug) at 95C 10min, chill on ice (I put on ice 10min), add mix with random hexanucleotides, Klenow, dNTPs, DIG-dUTP, etc, incubate at 37C 20 hours, stop reaction with EDTA.
3) Blot 1ul of DNA dilutions onto membrane along with 1ul of control DNA dilutions to determine labeling efficiency.
Technical support from Roche has suggested that I use a shorter probe (
I experimented with Dig labeling about 20 years ago and made my probes with PCR reaction. I found an increase in signal using sterile reagents (when possible) for hybridisation and detection.
No logic till now, but I find, amplicons are better labelled when cloned into T-vectors. I have no reason as of now, but I resorted to making probes from clones after experiencing the same problems. Also, I would suggest use of chemiluminescent substrates for detection, rather than colourimetric detection.
Thanks for the responses. I am now trying the PCR labeling with several different sets of primers to make different sized probes from 500-2,500 bp. Will see if I have any luck.
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