After PCR, One Band on 1.2% Agarose but 2 bands on 8M Urea, 8% PAGE - (Jun/28/2012 )

Hi,

I apologize ahead for the length of the post. I hope to include any necessary information. I am desperate to figure out what is going on. I perform PCR (see below for procedure), clean with QIAquick PCR clean-up kit and check purity on a 1.2% agarose gel (http://i1050.photobucket.com/albums/s410/jatracy/2012-06-27PCR800msedit.jpg) . I see one band that corresponds to the correct bp (310 bp). When I then set up my first control for DNase I footprints, I need to use a 8M, 8% PAGE. I prep my samples (see below) and then run it on the gel at 10W for 2.5 hours. On this denaturing gel, I now have a mysterious band that is much larger (around 1000-1200 bp in size compared to my 310 bp) than the PCR product used in the DNase I digest (http://i1050.photobucket.com/albums/s410/jatracy/2012-06-28Control1editscaled.jpg) . I am completely confused on what can be going on. Please help!!

PCR:
1 uL 15 ng/uL 2909 plasmid as template
3 uL each 10 mM Primers (one primer is labeled with a 5-FAM tag)
10 uL 5x Taq Master Mix from NEB
sterile water to 50 uL
*all solutions and reactions maintained on ice

Hot start (add samples once block reaches 85C), 2 min hold at 94C, 30 cycles: 30s 94C, 40s 56C, 30s 72C, hold at 4C until I remove (usually within 10 min of completion).

Control Rxn Setup:
(1) series of DNase dilutions: stock DNase is at 10mg/mL (35.33 U/uL) in 0.15 M NaCl. I create a 100x dilution to produce a 0.3533 U/uL solution in 50 mM MgCl2. This is my working DNase "stock". I then perform two dilutions: 1 uL stock into 9 uL 50 mM MgCl2; serial dilute 1 uL created dilution into 9 uL 50 mM MgCl2. I use these three dilutions to create a series of 12 dilutions. The buffer used to create the dilutions is 100 mM NaCl, 50 mM NaH2PO4, 5 mM MgCl2, 1 mM CaCl2, 2 mM DTT, 20% glycerol, 100 ug/mL BSA and 2 ug/mL Sheared Salmon Sperm (pH 8.0).
(2) Reaction tubes containing 150 ng 5-FAM labeled PCR product and Assay Buffer (same as buffer used in the DNase dilutions above minus the BSA and Sheared Salmon Sperm) are assembled. The total volume is 200 uL.
(3) 5 uL of DNase dilution is added to Assay Reactions, vortexed briefly, incubated at 37C for 2 min and reaction stopped by adding Stop Solution (16.125 mL 100%EtOH, 1.25 mL saturated ammonium acetate; 750 uL added to each reaction), vortexed for 5s and moved to -80C. Once all reactions complete, they are incubated for 25 min.
(4) They are then spun down and washed twice with 70% EtOH and dried for 5 min in Speed-Vac centrifuge.
(5) 5 uL 95% formamide is added, pellet resuspended (never completely dissolves), vortexed and spun briefly and incubated at 90C for 7 min. Samples are then moved to ice-water bath for 5 min and loaded directly onto denaturing gel.

Things I have tried:
- Using less template during PCR; just get less product
- Increasing annealing temp to 57C and 58C; still see above
- Ran a denaturing gel without the DNase treatment with the samples prepped both as above and by simply mixing DNA with formamide, heating for 7 min at 90C and loading onto gel. I still see this band appear only on the denaturing gel.

Thank you for any help and again apologize for the length of the post
jatracy

I realized I should describe the gels a little more. The agarose gel contains 100 bp NEB ladder, 4 lanes of PCR product prior to clean up and 10 lanes of the PCR products after cleaning. The bright band at the bottom of the gel is one of the tracking dyes in my mix (it lights up the same as SYBR Gold, which was used to stain the gel). You can see the excess primers in the first four lanes.

The Denaturing gel is imaged using Typhoon Trio Imager with a Blue laser and the 5-FAM label on the DNA sequence. Therefore the larger fragment also has this tag on it so must have the labeled primer in the sequence.