Bead Cleanup and dsDNA Quantification

Bead Cleanup

Dual selection to remove both high (gDNA) and low (primer/dimer) MW fragments

1. Mix 100ul beads per 200ul PCR reaction by pipetting (0.5:1 ratio).
2. Incubate 15 min at RT with mixing.
3. Incubate on magnet 10 min.
4. Remove 275ul solution to new tube while tube is still attached to magnet (large MW fragments remain on beads and are discarded).
5. Add 55ul beads per 275ul eluate (0.8:1 ratio, accounting for transferred solute).
6. Incubate 15 min at RT with mixing.
7. Incubate on magnet 10 min.
8. Remove and discard supernatant containing primer/dimer fragments. Keep the beads!
9. Wash twice with 200ul fresh 70% ethanol while tube is still attached to magnet (do not disturb the beads):
   Add 200ul 70% ethanol. Let sit 30 sec RT on magnet. Carefully remove ethanol wash. Repeat.
   *If using a large magnet, tilt on its side so ethanol covers beads and rock gently.
10. After removal of second wash, air dry beads 1 min.
11. Resuspend beads well in 35ul H2O.
12. Incubate 2 min at RT; place on magnet.
13. Run 2ul on a 1-2% TAE agarose gel to verify removal of unwanted fragments.

Measuring dsDNA Concentration with Qubit

1. Equilibrate all High Sensitivity dsDNA Qubit reagents to RT.
2. Set up Qubit assay tubes: sample number +2 for standards.
3. Prepare working solution in a plastic tube (200ul per sample + 200ul extra - no glass!). 1:200 reagent:buffer
4. Aliquot working solution to tubes; add standard or sample. Vortex 2-3 sec.
   200ul total per tube:
   For standard: 190ul working solution + 10ul standard
   For sample: 190-199ul working solution + 1-20ul sample
   
5. Incubate at RT at least 2 min.
6. Read on Qubit.