I isolated mouse genomic DNA and digested it with BglII at 37C overnight (5 ug DNA, 10x NEB Buffer 3.1, 20 U BglII, 0.3 ul RNAse cocktail, 0.3 ul BSA (20 mg/ml) in a total volume of 30 ul. I ran the DNA in a 0.8% agarose gel for 2 hr at 100 V along with a DIG-labeled MW ladder and unlabeled probe (797 bp). The probe was created using an initial PCR from genomic DNA followed by agarose purification and DIG-11-dUTP PCR. 

The gel runs reasonably well (see photo 1).

I treat the gel with 250 mM HCl for 20 min, followed by diH2O for 5 min, then 400 mM NaOH for 20 min, followed by diH2O for 5 min. I set up the transfer using + charged Nylon membrane and 20x SSC buffer. The next morning, I rinse the membrane with 2x SSC and UV crosslink for 5 min. I then dry the membrane and hybridize with Easy Hyb at 42C for 1 hr. I incubate in 20 ul of probe in 5 ml of Hyb buffer overnight at 42C. The following day, I wash in 1x maleic acid with 0.3% Tween-20 5 min twice and then block in Roche blocking reagent for 30 min. I incubate in 10 ml 1:5000 anti-DIG-AP for 30 min at room temp and wash twice with the wash buffer for 15 min. I set the membrane in detection buffer for 5 min and treat with CSPD. The images were collected after 5 min and 1 hr. The heavy band to the right of the ladder is the unlabeled probe control (1:100) and the bands I want to see are above and below the third marker band from the top. The lines above the wells were made with pencil, but the wells seem to react as well.

What should I try next?

Thanks!