Keegan, I doubt there is any organics-based DNA extraction method excluding phenol-chloroform or excluding any kit. However, if your sample size is small, and if your down-stream usage can tolerate some dirtyness, you could try something simple: make a suspension of your tissue/cells in a tris or phosphate based buffer (TBS/PBS) with detergent, digest with first RNAse and then add SDS to 1% and proteinase K overnight. Finally, precipitate with ehanol. You will retain some degraded proteins in the preparation, and might struggle to get the final pellet into solution. You have to titrate the amount of RNase/PrK for the amount of cells/tissue lysate.

I can suggest a way to optimize for yourself, though! DNA binds calcium phosphate gel (classically known as hydroxyapatite) at 500mM NaCl (to do a pilot study, take a cell of interest, or closest to your interest; say, HEK293/NIH3T3; make the suspension to a known concentration, e.g., 1M cells/ml). 2. Wash a bulk of hydroxyapatite (not expensive at all) in the same buffer, aliquot the hydroxyapatite suspension (say, 100ul) into each tube. 3. Pippet varying amount of lysates into the hydroxyapatite aliquots in replicates (that is, each amount of lysate to be taken in duplicate/triplicate). 4. Bind the lysate to hydroxyapatite in cold room for 1-2hr (rotate). 5. Wash the beads in the binding buffer 2-3 times. 6. Elute the DNA twice in, say, 200ul 0.5M phosphaste buffer (100ul twice). Precipitate the DNA. If using eukaryote DNA, you should expect to precipitate the bulk of histones as well.

By varying the amount of hydroxyapatite and the lysate, you can determine the effective ratio within a couple days. In another couple days, you can repeat the procedure with different cell types/tissues. In a week's time, thus, you will have your own protocol and Kit. You can make the DNA preparation cleaner by increasing NaCl concn in the binding/washing buffer. Cool?

Forgot to mention: any cell lysis buffer much contain some detergent when mechanical shearing is avoided. So, do not try to lyse the cell suspension with syringe or sonication. Have Triton or Igepal at, say, 1% (may try sod. deoxycholate also; see which combination works better without hampering binding to hydroxyapatite). The good thing is, hydroxyapatite is compatible with most detergents commonly used in protein purification.

There was a BioTechniques paper years ago on how to reuse spin column filters from kits (which give fairly pure DNA), since the reagents could be made in the lab.

These are both very interesting options that I'll have to look into, thanks for the advice!

Gary, we have a few spin columns spare from old kits laying around - if we can re-use them without risking cross-contamination then that would be great!

Anil, thanks so much for all the detail. What kinds of tissues are you normally working with for your lysis step? We have some very tough samples, and I don't know if we can sufficiently interrupt the cells with an overnight digestion - in the past, we've only ever had luck with mechanical approaches like bead-beating. If I was to bust the samples up mechanically and reduce the detergent concentration, do you think that might work?

Keegan, I pretty much work on cultured cells. But I know there are some automated, easy-to-use tissue dissociators, like https://www.miltenyibiotec.com/en/products-and-services/macs-sample-preparation/sample-dissociation/gentlemacs-dissociator/gentlemacs-dissociator.aspx, that you may want to consider. Reducing the detergent concn may not be the option in my opinion. The pilot study that I proposed earlier would best suit cultured cells, so I concede that there has to be lots of ground work to do. I shall let you know if I get some better thoughts.

Hi Anil, thanks for all the help just the same! It may be that we can work out a fusion of techniques that will suit our samples, so I'll definitely keep this outline in mind during the next stages of work!