Joel Huberman's DNA isolation procedure with modifications by Bonny Brewer
1. Kill cells at mid log phase by adding Na Azide to 0.1%. Immediately chill culture by adding it to frozen 0.2 M Na EDTA. (40 ml of EDTA/200 ml of culture) Shake as you would for a martini to chill the cells. Spin 6K 5 min to pellet cells.
2. Wash cells in ice-cold water. Transfer cells to a conical-bottom 50 ml Falcon 2098 tube. (At this point you may freeze the cells as a "dry" pellet at -20.)
3. Resuspend the cells at 1.5 to 2 X 109/ml in ice-cold Nuclear Isolation Buffer (NIB = 17% glycerol, 50 mM MOPS buffer, 150 mM potassium acetate, 2 mM magnesium chloride, 0.5 mM spermidine, and 0.15 mM spermine; pH is adjusted to 7.2 after all ingredients are dissolved).
4. Mix cell suspension with an equal volume of acid washed glass beads (0.45 to 0.52 mm diameter). Keep on ice.
5. Vortex at maximum speed on a healthy vortexer for 30 sec periods. Chill on ice for at least 30 sec between episodes of vortexing. Repeat the vortexing 10 to 15 times or until more than 90% of the cells have been broken (monitor by checking for the fraction of ghosts in the phase contrast scope). Breakage is most efficient when the entire contents of the tube are lifted by the swirling action. I also find that erring on the side of having too many beads aids in breakage.
6. Remove the supernatant with a glass pasteur pipette or a blue pipeteman tip--stick the tip through the beads to the bottom of the tube and suck up the liquid. It doesn't matter if a few beads get transferred. Rinse the beads twice with 1.5 volumes of fresh NIB. Pool and spin at 8 K in an SS-34 rotor for 20 min.
7. Resuspend pellet of nuclei, ghosts, and unbroken cells at a final concentration of 2 X 109 cell equivalents/ml in 50 mM Tris, 50 mM EDTA, 100 mM NaCl, pH 8.
8. Add Sarkosyl to achieve a final concentration of 1.5%. Mix gently.
9. Add solid Proteinase K to a final concentration of 300 microgram/ml. Incubate at 37o for 1 hr.
10. Centrifuge (cold) at 5 K for 5 min to pellet cells and ghosts. The supernatant should have only minor turbidity.
11. For each ml of supernatant soution, add 1.05 g of CsCl. (I aim for a supernatant volume of 4.0 ml (or 4.08 g) plus 4.2 g of CsCl. This volume will fill one VTi65 heat seal tube.) Encourage the CsCl to dissolve by gentle mixing. Don't mix vigorously or foaming will result.
12. After the CsCl has dissolved, measure the volume and add 0.025 volumes of a 5 mg/ml stock solution in water of Hoechst 33258 dye. Mix.
13. Transfer to 5 ml heat-seal tubes. Fill the tube to the top with additional "dummy" solution. ("dummy" is the same solution--NaCl, Tris, EDTA, CsCl, Sarkosyl, Hoechst--minus the yeast DNA).
14. Spin in a VTi65 or VTi65.2 rotor at 55K and 20o for 18-24 hr.
15. Visualize the DNA bands under long wave UV light. Theoretically there are three bands. The top, diffuse band is mitochondrial DNA. The prominant band is chromosomal DNA and the faint band immediately below is the rDNA band. (Hoechst separates DNA based on GC content.) You may notice a band of particulate junk between the nuclear and mitochondrial bands. Stay away from this stuff. Remove the DNA by side puncture using a 16 gauge needle. Measure the volume in the syringe before dispensing it into a 15 ml corex tube. (I usally remove the needle before dispensing it to reduce the possibility of added shear.)
16. Add an equal volume of 5:1 isopropanol:H2O solution. Swirl the tube to mix the contents (I don't use the vortexer because of worry about shear). After the phases separate, remove the alcohol phase with a pasteur pipette. I let the phases settle out in the drawn out tip of the pipette and use it like a separatory funnel to minimize the loss of any aqueous phase. Repeat the isopropanol step twice more.
17. Add 3 volumes of cold 70% ethanol slowly, to the side of the tube. With the ethanol floating on the CsCl layer begin mixing the phases with a single quick swirling motion. The DNA should begin to come out of solution at the interface as a fibrous network. Continue the single swirls until the phases are mixed and the DNA has fallen out of solution. The DNA can be removed by touching the DNA clot with the end of a pasteur pipette that has been closed by melting it in a flame. The DNA will stick to the glass and can be submerged in fresh 70% ethanol to rinse the clot and then teased from the end of the glass tip onto the side of a microfuge tube. Don't let the clot dry out completely before adding TE to redisolve the DNA. If you are looking at plasmid replication intermediates, spin the ethanol-CsCl solution 20 min at 8 K. Rinse the tube with fresh 70% ethanol, air dry the tube and resuspend any DNA in TE. The two DNA solutions can then be pooled. For a 500 ml culture I would resuspend the DNA in a final of about 400 microliters. It may take the clot several days to go into solution. If you are worried about losing small bubbles you might want to add NaCl to 50 mM after the DNA has begun going into solution.
18. To look at single copy sequences I usually cut about 1 microgram of the nuclear DNA in 0.12 to 0.15 ml of buffer with 4-5 microliters of enzyme for 5 hours. Digestion may be complete with less enzyme or with shorter incubations but since it is likely to vary with the enzyme I haven't strayed much from this plan. The DNA is precipitated by the addition of 2 volumes of absolute ethanol containing 0.5 M K-acetate, chilling for 15 min at -70o, and spinning 15 min in a microfuge. After rinsing the pellet in 70% ethanol and drying the tube I resuspend the pellet directly in loading buffer.
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