We use a mixture of phenol, chloroform and isoamyl alcohol to extract protein/lipid contaminants from mammalian genomic DNA. The phenol is pre-equilibrated to a basic pH with a saturated Tris solution. One part of the lower phase (organic) of the phenol is mixed with one part of a 24:1 mixture of chloroform and isoamyl alcohol. If this solution is allowed to sit, an aqueous layer forms on top. Presumably, this represents water and Tris that equilibrated in the phenol layer but are now forced out by the increased hydrophobicity of the solution.
My contention is that it is improper to use the resulting organic phase alone to extraxt DNA. Better, would be to shake the solution prior to use so that the aqueous solution could be co-transferred. My rationale is that the purpose of adding the aqueous tris to the phenol is to increase the pH of the extraction mixture so that depurination of the DNA bases does not occur.
Preface: I know little about the biological end of biochemistry.
That said, I really don’t see what the problem is. Perhaps I’m missing something, but this appears very simple.
It sounds like the entire point of this step is to remove the aqueous phase and the impurities within (including, perhaps excess tris?) and that mixing the two will negate the entire purpose of this step of the extraction.
If there is a pH problem in a subsequent step, it would be better to add pure tris (or whatever) just before that point and get rid of the mix of impurities in the aqueous phase now. Is there some reason why it’s difficult to measure pH, so that you could say for sure that it was a problem? Are you troubleshooting this procedure because of depurination of the final product?
In general, if your desired product isn’t in one of the phases, throw that phase away… that’s the whole point of doing an extraction.
I think you are correct here. The recipes call for phenol/chloroform/isoamyl alcohol (25:24:1). If you let the (25:24:1) mixture separate, neither phase will have that composition. If the proper method of preparing the extraction buffer involved collecting only the lower phase of the phenol/chloroform/isoamyl alcohol mix, that process would be included as a step in the buffer preparation protocol. It is not.
Tim is favoring what would probably be the right approach for an oganic chemist, but proteins are remarkably picky about precisely how they are denatured and removed from a solution.
Thanks for your reply, (Tim) (Say, those parentheses are nifty, but I hope you won’t take offence if non-touch typing me drops them for the rest of your tenure on the boards:)). I think I need to explain the OP a bit. You see, the Tris solution is added to the phenol before any other mixing takes place. The Tris solution used is a highly pure “molecular biology” grade so I doubt it contributes much in the way of impurities. It is added to phenol prior to use in DNA extractions to raise the pH of the pure phenol (phenol is a weak acid, and low pH solutions depurinate DNA creating a more-easily degradable molecule). My understanding is that much of the water and Tris dissolves in the pure phenol because although is a fairly large organic molecule, it has a polar hydroxyl group capable of hydrogen bonding.
Tris-equilibrated phenol alone is commonly used in DNA extractions. Phenol is a great solvent for proteins (don’t get it on your hand–ouch!) but works less well for lipids. The point of adding the chloroform and isoamyl alcohol to the mix, IMHO, is to better extract lipids from the tissue mixture. Thus, I do not believe these additions are aimed at he removal of the dissolved Tris (or other impurities)from the phenol, although this occurs as a side effect.
My contention is that it is a potentially important side effect, however, since the entire point of adding the Tris to the phenol was to create a DNA-safe environment.
This is sort of what I was thinking. BTW, Squink, I’m not trying to clarify a protocol ambiguity. I’m simply trying to correct a know-it-all technician in our lab who insists that allowing the phases to separate before use is the way to go (he even spins the shit in our cytocentrifuge to layer it out faster which invariably coats the inside of said centrifuge with phenol). Personally, when I do these extractions (and I make virtually any excuse to get out of doing them, they suck, bad), I add the phenol and chloroform/isoamyl separately to the extraction tube. I’ll follow up on your suggestion to find protocols, however unless I find one that’s awfully clear and authorative, I doubt our mule-headed technician will stop stinking up the centrifuge.
My point was that the protocol would never leave such a complicated step in buffer preparation to chance. The protocols were written so that undegrads and high school students can do them without thinking. They are very explicit. A simple reference to a 25 : 24 : 1 mixture of phenol : chloroform : isoamyl alcohol mixture refers to the complete mixture, not some mystery mix that you get by sucking off half the water and much of the Tris.
You do it the way I did it, and the way the guy who taught me to do it did it, and there’s just not a lot of sense in thinking that it “should” be done in some peculiar way that’s not described in procedures that have been handed down since the dawn of time, (or at least since the early seventies).
The word ‘extract’ in the first sentence made me think the step you were referring to was intended to be an extraction. Yeah, that’s the ticket.
Just to clarify one minor point: I wasn’t claiming the tris contained impurities, just that any polar impurities would be concentrated in the aqueous layer. It seems these aren’t relevant and the amount of tris is. I understand.
I’m still curious, though. It sounds like this procedure has been done with both removal of the layer and without, do you know offhand if and how the final results were different? Or is that not easily quantifiable or directly comparable?
Yes.
It’s an early step in purification that needs to be able to handle large variations in the chemical composition of the cells being extracted. It’d take a large effort to optimize the extraction for a given cell type, and in most cases here, the percent yield isn’t as important as following the same reprducible procedure time after time.
That’s a tough call. We use the extracted DNA to do very messy southern blots or messy PCR for transgene screening. High quality DNA is not particularly necessary for either procedure. I notice no difference in the results obtained using DNA from either extraction procedure. However, I’m certain a more sensitive assay would reveal some DNA degradation in the incorrectly extracted sample.
What bugs me is the needless centrifuge contamination and the tenacity with which the tech in my lab grips his ignorance.
The first is that for PCR, phenol extraction is virtually never necessary, and often adds to your problems. In fact, I do no purification beyond the initial lysis/digestion step (NaCl, Tris, EDTA, detergent, Proteinase K), just leave the samples in lysis buffer overnight so that the Proteinase eats itself up. Make sure that your detergent is Sarcosyl or Triton, not SDS, since SDS inhibits PCR. You may have to re-optimize your PCR (esp. Mg concentration), but it will soon save you SO much time. This works fine for PCRs up to about 2-3 kb, if yours is longer, you will need to purify the DNA. Note: more is less, I use 0.3ul of DNA sample out of a 500ul tail sample and get reliable genotyping, everytime. More tends not to work as well, presumably because of the increased amount of detergent added to the mix.
For mammalian Southerns, I find a simple salt precipitation of contaminants and some ethanol washes works better than phenol precipitation - I find that the phenol is hard to completely remove from genomic preps, and it inhibits the restriction enzymes. Email me if you are interested in detailed protocols we use in my lab.
As to the phenol (your original question), it is best to leave a layer of aqueous Tris on top, to maintian the Ph. It is generally desirable to let these layers settle out, because you don’t want your aqueous DNA sample to be diluted by the excess aqueous Tris solution that is still mixed in to the phenol/chloroform/isoamyl alcohol. However, this doesn’t matter much, especially if you are going to precipitate the DNA anyway. And tell your tech to just leave the damn stuff in the fridge overnight, it will settle fine. Contaiminating the centrifuge is a serious hazard - you know that phenol will raise blisters on skin contact, but did you know that ALL three ingredients are liver-damaging agents when inhaled. (Can you tell I’m chemical safety officer for my lab?)
Sorry I’m arriving late to this discussion, but I only “lurk” at work, and post only from home.
choosybeggar, I have done literally thousands of these extractions (I know, lucky me!). They were all for “non-messy” Southern blots, not PCR (I considered “Blotmeister” as a username). I pretty much did it the way you apparently do. Two references I checked, including Current Protocols in Molecular Biology, make no mention of removing the aqueous layer from the 25:24:1 phenol:chloroform:isoamyl alcohol, either. As others have said, if it was necessary it would be in the protocols.
In my experience, it is next to impossible to pipet the Tris saturated phenol without taking at least a little of the Tris layer along with it. So you add that to an equal volume of 24:1 and mix (vortex, even). You end up with 25:24:1, with tiny droplets of aqueous Tris dispersed throughout. I (and you, apparently) just pipet that out and go from there. Your compulsive lab-mate insists on spinning this down so that the Tris is now on top, and then uses the lower layer? This strikes me as an extra step that wastes time as well as, (as mischievous points out), creates a genuine health and safety hazard. The only drawback to “our” way is that you are slightly diluting your aqueous DNA layer.
In my informed opinion, either way should work just as well. It seems to me the burden would be on the tech to prove that their longer, more dangerous procedure actually produces superior results. Is the, er, esteemed colleague using 1.5 ml microcentrifuge tubes? If they absolutely must, they should be able to spin these down without contaminating the whole microfuge, but he/she needs to not fill them up to the top.
And, as mischievous also points out, by far the easiest/safest way to separate out the aqueous is to just leave it at 4C overnight. In fact, Current Protocols says that this mixture can be stored for months like this (just wrap the bottle in foil).
Actually, I have a related question: when you store phenol or the phenol/chloroform/isoamyl alchol mix at 4C, it eventually goes bad. The indicator you are suppossed to use to know if it is bad is the color - clear is good, pinkish or brownish is bad. Now, in my laziness, I almost never make up the poison stuff (and almost never use it), so when I do get around to using it, it is almost always pink. And it doesn’t seem to make a damn bit of difference. Has anyone else had this experience?
