Performing Minipreps with Homebrew Buffers

The alkaline-lysis miniprep is a critical tool in the arsenal of a molecular biologist. It allows one to rapidly isolate only plasmid DNA from a bacterial cell by leveraging the increased resilience of this (usually supercoiled) form of DNA against highly basic conditions.

Minipreps are routinely performed to isolate plasmids that serve as substrates for further assembly work, for PCR amplification of specific gene segments, for direct application in other species or strains, or simply for archival uses (DNA can be easier and cheaper to store than the cells containing it).

However, minipreps are usually done today with kits, using convenient but expensive sets of buffers and binding-columns. Not only are these harder to get as a DIYbiologist, but they actually provide lower yields and may cause shearing or nicking of DNA.

Thankfully, miniprep buffers are not so complicated that they can’t be made at home, provided you can get the required chemicals (some of which are surely replaceable). Preparing buffers takes a while, but you’ll be making volumes of 100mls each; enough for hundreds of minipreps. Here’s how.

Warning Preamble

You’ll be using strong mineral acids such as Hydrochloric Acid (HCl) and strong mineral bases such as Sodium Hydroxide (NaOH) if you want to make the below buffers DIY style. Wear gloves at all times and consider goggles when handling these chemicals. Do not measure chemicals (particularly NaOH) with/into metal (particularly aluminium), as runaway reactions can occur. Equilibriate buffers slowly and stir constantly to prevent overheating due to acid/base reactions. (See footnote 2)

Ingredients you’ll need:

  • A pH meter to prepare the buffers, calibrated. Handheld pool testing pH meters are probably the easiest and cheapest way to go: one of these is what I use in the lab.
  • A centrifuge to actually perform the miniprep. You might be able to forego this by binding DNA to glass or silica, but I haven’t tried. I used my dremelfuge, though I had to leave it to cool off between extended spin cycles.
  • You’ll need pipettes accurate enough to enable precise buffer volumes; you may be able to do this with a 1ml glass pipette if you’re very good, but I suggest investing in some micropipettes.
  • Tris: A common alkaline buffer, useful for getting accurate pH values around 8-10.5 or so. Despite good safety profile, probably the hardest chemical to find on this list. Can likely be replaced with another buffer but I have yet to test this.
  • EDTA: A chelating agent used to protect DNA, and a mild acid. If you can’t get any, you can do without it but your DNA will not be very stable due to the action of ubiquitous nucleases.
  • Acetic Acid: You really won’t be able to do this with only 5% acetic acid, known commonly as vinegar. Ideally, 50% or better is needed. However, if you can’t get strong acetic acid, you can prepare stronger acid from weak solutions (vinegar) by boiling away the water, or freezing out the acid.
  • HCl: You could probably forego this in favour of acetic acid, if you can’t find it. However, HCl is commonly available as “Spirits of Salt” in hardware stores, and can be easily found online.
  • NaOH: Sodium Hydroxide, or Caustic Soda. Easily available in hardware stores or supermarkets. Surprisingly dangerous for a household chemical: Do not allow to come in contact with aluminium or other metals, or runaway heating/boiling/burning can occur, in addition to production of explosive H2 gas.
  • Sodium Lauryl/Laureth Sulphate (SDS or SLES): Although traditionally sodium lauryl sulphate is used (also known as sodium dodecyl sulphate in most labs), I successfully used sodium laureth sulphate, a closely related detergent found in many household products and more easily found online.
  • Sodium Acetate: Although you can prepare this easily from acetic acid and sodium hydroxide, the amounts needed may merit separate purchase. If you can’t find “pure” anhydrous sodium acetate (as well you should), you can get sodium acetate trihydrate from uncoloured reusable heat-packs, the sort activated by clicking a piece of metal and recharged by boiling. The contents should be fairly pure sodium acetate trihydrate. Use in its solid form, not the highly energetic liquid form!
  • Lysozyme: This enzyme, used to chew away bacterial cell walls, can either be purchased inexpensively from brewshops (the brand I use is called “Maltostop” and was bought from, or prepared in-lab by combining 10mls of egg white with 30mls 40% vodka, leaving for 3hrs, diluting 50%, filtering, and drying out under conditions that don’t destroy the enzyme (probably impossible without dialysis or a vacuum pump).
  • Isopropanol or Ethanol: Given how difficult it is to find pure ethanol in Ireland, I use Isopropanol. The two alcohols yield good results, but you need less IPA than EtOH. The disadvantage of IPA is that the precipitated DNA forms a clear and glassy pellet, whereas EtOH results in a clearly visible white pellet. You can make do with less-than-pure alcohol, but it’s very worth investing in pure: The volumes needed to precipitate DNA spiral the weaker the alcohol gets. Below 70%, forget about it.
  • Deionised Water: Don’t skimp and use tapwater. Go get deionised water, or you risk the entire thing failing. I used the kind you buy cheaply in supermarkets for old car batteries or for filling steam irons, and it worked fine. Distilled water is not necessarily any good, either; if prepared/stored in/via metal containers, it may carry metal ions that help chew up DNA. Wherever water is mentioned or implied, assume it’s deionised water.

Buffers to Prepare:

Resuspension Buffer: 5mg/ml Lysozyme, 25% w/v Sucrose, 25 mM (0.303g/100ml) Tris, 10 mM (2.92g/100ml)  EDTA, slowly and patiently adjusted to pH 8 with HCl. I’d highly recommend that you prepare this fresh and then aliquot into small volumes and freeze until use, to keep the lysozyme from degrading. Alternatively, make up 2x Tris/EDTA and add freshly prepared 50% Sucrose solution with 10mg/ml Lysozyme solution right before use.

Lysis Buffer: 0.8g NaOH + 1% v/v SDS/SLES in 100ml deionised water.

Neutralisation Buffer: 29.6g Sodium Acetate (NaAc) dissolved in 30ml 50% acetic acid, slowly adjusted to pH 4.8 with strong sodium hydroxide solution (stirring constantly) then raised to 100ml with deionised water. See footnote 1.

Washing Buffer:  25 mM (0.303g/100ml) Tris, 10 mM (2.92g/100ml)  EDTA, 0.3M (2.46g/100ml) Sodium Acetate in 100mls (You can equally just make 3M Sodium Acetate solution (24.6g/100ml) and add to nine parts of whatever solution contains your DNA).

Tris-HCl-EDTA: Unnecessary but highly recommended; TE buffer helps to protect your DNA from nucleases. Prepare with 25 mM (0.303g/100ml) Tris, 10 mM (2.92g/100ml)  EDTA, adjusted to pH 8.5 with HCl solution before being raised to 100ml (i.e. prepare as a 2x solution, adjust pH, then bring to 1x).

Procedure for Plasmid Extraction

  1. Grow your cells overnight in small volumes, shaken if possible to maximise aeration and culture growth. If your plasmid requires antibiotics, be sure to add enough to maintain the plasmid throughout growth, and optionally rinse cells (by centrifuging to remove broth, resuspending in fresh broth, and repeating this once again) before inoculating the culture to wash away extracellular antibiotic-degrading enzymes.
  2. For a high-copy plasmid, take 1.5-3mls of cells from the overnight culture (leave some cells from which to grow more plasmid if you don’t already have stocks!), and centrifuge to isolate the cells as a pellet.
  3. Resuspend cells in 100uL of Resuspension Buffer. For E.coli, leave to sit at room temperature for 5 minutes. For B.subtilis, leave for 30 minutes at 37C. Cells should look the same as usual; cloudy and prone to settling if left for too long.
  4. Add 200uL Lysis Buffer and immediately cap and invert the tube several times to mix. Don’t vortex, pipette up-and-down or otherwise violently mix the sample, or you may shear genomic DNA, causing contamination of your plasmid extract. You should see the samples clarify as cells burst open in the detergent. Leave samples for 1 minute. Don’t leave for longer than this.
  5. Add 150uL Neutralisation Buffer and mix as before by capping and inverting repeatedly. You should see a white precipitate form: for high-density E.coli this is often a dramatic white clot, but it is often more subtle. This is a combination of denatured protein and genomic DNA.
  6. Centrifuge your samples at maximum speed for 5 minutes to form a hard pellet of denatured cell contents.
  7. Remove the supernatant (remaining liquid after pelleting of precipitate) to another centrifuge tube. This contains the still-dissolved plasmid DNA, which survived denaturation and precipitation by virtue of its supercoiling.
  8. To this, add 0.5ml chilled IPA, mix by repeated inversion, and leave to precipitate for 10 minutes.
  9. Centrifuge tubes at maximum speed for 10 minutes. After this, a small glassy pellet should be barely visible at the bottom of the tubes; black marker may help make it more visible. This is not pure DNA; it will contain cellular RNAs in addition to whatever plasmids may have been extracted. Without RNAse, it is difficult to separate RNA from DNA without gel extraction.
  10. Remove as much of the supernatant as you can without directly disturbing the pellet, leave to dry (but not completely) for 5 minutes, and add 0.7ml DNA Washing Buffer. Redissolve DNA by shaking or inverting, possibly after warming the tube in hot water. The pellet need not be entirely redissolved; the aim is merely to resuspend it somewhat to help rinse away soluble salts and remaining miniprep buffers.
  11. Re-precipitate the DNA with 0.7ml IPA for another 10 minutes, followed by another 10 minute centrifuge cycle.
  12. Remove as much of the supernatant as possible and leave tubes open for 10 minutes to allow IPA to evaporate. If the trace remaining bits of buffer don’t halve in this time due to evaporation, leave for another 10 minutes or consider sitting tubes in hot (but not boiling) water. Don’t allow samples to fully dry out; once most of the liquid is gone, it is likely that all IPA is evaporated.
  13. Resuspend and dissolve the pellet in a desired amount of hot (but not boiling) TE buffer. 50uL-100uL is suggested. In TE buffer, DNA should be stable for weeks or months at 4C. In deionised water without EDTA or buffering, you may be better off freezing samples if they won’t be used by the end of the week.
  14. You can visualise your samples on a gel to see if the miniprep worked. If you have not cut the DNA with a restriction enzyme before doing so, you may see two smaller-than-expected bands in your gel, corresponding to supercoiled DNA and DNA which has been “relaxed” by accidental nicking of one strand during processing. If you do cut your DNA, you’ll either see only one band corresponding to the correct size, or simply an additional band corresponding to the correct size (if digestion was incomplete).


  1. I did this the other way around; NaAc dissolving in water and attempting to bring to 4.8 with acetic acid. It took so much acid that I ended up having to double my volumes, and I ran out of acetic acid and had to finish the job with HCl. Dissolve in Acid and equilibriate carefully with base instead. It’s awkward and time-consuming: be careful and patient, because you can crawl up by decimal-points of a pH for ages and then suddenly crash past the target by several pH points as the buffering capacity of the solution is exceeded. Slow, carefully and patiently is key.
  2. I made the mistake of using a little bit of aluminium foil as my weigh-boat. While looking away for a minute, I heard a fizzing/popping noise and looked back; the sodium hydroxide had gathered air moisture (it’s highly hygroscopic) and was dissolving itself on top of the aluminium. The fizzing was coming from the production of hydrogen gas. I threw it into a large volume of water and removed the foil when it was clear of NaOH. The volumes weren’t big enough in this case to cause much worry, though droplets of concentrated NaOH were being spattered about; if I wasn’t wearing gloves I’d have lots of tiny burns. I happen to know someone who nearly blinded a friend of his while playing with NaOH and aluminium foil when the container melted and the solution flash-boiled.