1. Run the restriction digest and the appropriate size markers on an agarose gel with ethidium bromide. Leave at least 1-2 wells between samples. Run the gel until the DNA bands are well separated as visualized on the UV box. Use fresh running buffer to minimize plasmid contamination.

2. Make sure the UV box is well cleaned or put a Saran wrap between the UV box and the gel. Use a clean razor blade or scalpel, cut a slit just ahead of the band of interest. Cut a slit just behind the band of interest.

3. Cut two small pieces of NA-45 paper to the width of the band. Using a forceps carefully insert a NA-45 paper into the slit ahead of the band and a NA-45 paper into the slit behind the band.

4. Make sure the NA-45 papers will not float up once placed in the running tank. Place the gel in the running tank and run the gel until the band of interest has moved out of the gel and onto the NA-45 paper ahead of the band. Other bands of higher molecular weight should be stopped by the NA-45 paper behind the band of interest. Estimate how far the band has moved by mark the position of the dye. Confirm by visualize under the UV light.

5. Remove the NA-45 paper ahead of the band (discard the one behind the band), rinse in water or TE buffer and place in an eppendorf tube. Add 100-300 ml TE, and 0.2 volume 5M NaCl. Submerge the entire strip.

6. Place at 65 °C for 0.5-1 hour. Make sure the NA-45 paper is fully submerged. DNA will be eluted from the NA-45 paper.

7. Remove elution to a clean eppendorf tube. Cool on ice.

8. Add equal volume of phenol:choloroform:isoamyl alchahol (25:24:1). Vortex, spin at 12,000 rpm for 10 minutes. Transfer the aqueous phase to a clean eppendorf tube.

9. Add 2 volumes of ethanol and precipitate the DNA at -20 °C for 1 hour.

10. Pellet the DNA by centrifugation at 12,000 rpm for 20 minutes at 4 °C. Discard the supernatant.

11. Wash the pellet with cold 70% ethanol. Be careful not to disturb the pellet.

12. Air dry and resuspend DNA in 10-50 ml TE.