Phenol-chloroform extraction

Phenol-chloroform extraction is a liquid-liquid extraction technique in biochemistry and molecular biology for purifying DNA contaminated by histones and other proteins. Equal volumes of a phenol:chloroform mixture and the aqueous DNA sample are mixed, forming a biphasic mixture. The proteins will partition into the organic phase while the DNA (as well as other contaminants such as salts, sugars, etc.) remain in the aqueous phase. This is usually repeated at least once and often more, depending on the requirements of the downstream processes, and then followed by precipitation with ethanol. Phenol and chloroform are both hazardous and inconvenient materials, and the extraction is laborious, so in recent years many many companies now offer many alternative ways to isolate DNA.

Reagents

• Phenol. The phenol used for biochemistry comes as a water-saturated solution with Tris buffer, as a Tris-buffered 50% phenol, 50% chloroform solution, or as a Tris-buffered 50% phenol, 48% chloroform, 2% isoamyl alcohol solution (sometimes called simply "25:24:1"). Phenol is naturally somewhat water-soluble, and gives a 'fuzzy' interface that is sharpened by the presence of chloroform. The isoamyl alcohol reduces foam, which is a problem with phenol:chloroform. Most solutions also have an antioxidant, as oxidized phenol will damage the DNA. Pure phenol crystals are no longer common. These had to be equilibrated into the buffer and then melted and dissolved, with care taken to avoid inhalation of the fumes or fine aerosolized powders.
• Chloroform. The chloroform will be stabilized with small quantities of amylene or ethanol because storage of pure chloroform solutions in oxygen and ultraviolet light will produce phosgene gas. Chloroform sometimes also comes as a 96% chloroform, 4% isoamyl alcohol solution that can be mixed with an equal volume of phenol to make 25:24:1.
• Isoamyl alcohol. Whether to purchase isoamyl alcohol separately or mixed in with the phenol and chloroform stocks is largely an individual choice, although it is sensible to coordinate with the rest of the lab so reagents can be shared. Some protocols do not require it at all.

Protocol

Additional recommended knowledge

Weighing the right way

What is the Correct Way to Check Repeatability in Balances?

Safe Weighing Range Ensures Accurate Results

Specific protocols vary between labs and even individual workers. A sample is given below. Note that a "volume" means an amount equal to the volume of the original DNA sample, e.g., 100μL is one volume for a 100μL sample.

1. Put 100-700μL of sample into a 1.5mL microcentrifuge tube.
   • Use water to dilute samples smaller than 100μL. The aqueous phase must be thick enough to see and remove, which is difficult for volumes less than 100μL. After ethanol precipitation, the DNA pellet can be resuspended at any desired concentration.
   • Divide samples larger than 700μL into multiple tubes. A sample larger than 700μL will not fit in the tube with a volume of phenol. If this is inconvenient, protocols exist that use tubes as large as 50mL, with special precautions taken to ensure even mixing.
2. Add an equal volume of phenol to the tube.
   • Phenol:chloroform:isoamyl alcohol will give a sharper interface, and 25:24:1 will show less foam.
   • Although pipettes and micropipette tips are usually resistant, phenol is known to attack polycarbonate plastic (clear and hard). For phenol:chloroform mixtures or for chloroform, glass pipettes should be used, or micropipettors exclusively, as the chloroform is usually able to attack plastic pipettes. In general, work as quickly as possible without sacrificing accuracy.
3. Vortex vigorously to mix the phases.
   • Small plasmids can withstand vigorous vortexing. It is even safe, although usually unnecessary, to hold two tubes together in the vortex cup so they collide violently during the vortexing. However, as the length of the DNA increases, so does the risk of shearing. Inverting 5-10 times by hand is a gentler method, and slowly turning the tube on a motorized rotisserie-style mixer for 30 min is gentle enough for DNA of any length. If necessary, try each method on a test sample and check for shearing on a gel.
4. Centrifuge at top speed for 1–2 min to separate the phases.
   • If the interface is disturbed, try letting the rotor decelerate without braking.
5. Remove the aqueous phase to a new tube.
   • The aqueous phase is usually the upper one. However, the difference in density is minute, and salts will invert the phases. Phase inversions are usually obvious because the organic phase is colored by the antioxidant. When the mixture is of water above 25:24:1 or phenol:chloroform, inversion is more difficult.
   • For valuable samples, the loss of DNA into the organic phase can be reduced by adding a second volume of water, mixing, centrifuging, and removing again. The second volume is combined with the first if space permits, or carried through the procedure in a separate tube. The added trouble of these extra tubes outweighs the benefits of increased yield for less valuable samples.
   • Removing the lower phase first can make the upper phase easier to remove.
6. Extract with a volume of phenol:chloroform.
   • Phenol is used for the first extraction because it removes proteins well. However, especially after repeated extractions, the aqueous phase will take in some phenol, and this step removes it to avoid interference in procedures downstream. Using phenol:chloroform or 25:24:1 instead of phenol for the first extraction reduces the amount of contamination, but many workers will extract again with phenol:chloroform.
   • Optionally, extract once more with chloroform. This is almost never necessary, but will ensure that no phenol whatsoever remains.
7. Ethanol-precipitate the nucleic acid