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A Menagerie of Mini Prep Methods

Over the years there have been a multitude of methods – ranging from the easy to the downright painful – developed for the preparation of plasmid DNA, but I suspect most of us have only used one , or maybe two, of these in our careers.

At Bitesize Bio, (part of) our life’s mission is to help you build your molecular biology arsenal so in this article I’ll brief you on the different plasmid miniprep methods available. If you have (or have had) a go at any of these, or if I’ve there’s a method I’ve missed, let us know in a comment.

Alkaline Lysis Method (reference here)

This method is based on the lysis and differential denaturation of chromosomal and plasmid DNA in order to separate the two (see here for more detail)

The lysis is performed using a NaOH solution containing SDS. During this step, both chromosomal and plasmid DNA are denatured, but upon neutralization using a high molarity sodium acetate solution, only the plasmid DNA is able to renature and remain soluble whereas the chromosomal DNA forms a complex with cellular proteins, potassium and SDS which is then removed by centrifugation. The plasmid is then isolated by precipitation using isopropanol.

This method creates plasmid generally suitable for most applications, although sequencing would need an extra phenol cleanup to make sure there is no extra soup contaminating your reaction.

Reagents required:

STE (sucrose/Tris/EDTA) solution (optional wash buffer):
8% (w/v) sucrose, 50 mM Tris-HCl (pH 8.0), 50 mM EDTA (pH 8.0). Autoclave and store at 4°C.

GTE (glucose/tris/EDTA) solution:
50 mM glucose, 25 mM Tris-HCl (pH 8.0), 10 mM EDTA (pH 8.0). Autoclave and store at 4°C.

Alkaline SDS solution: 0.2 N NaOH, 1% (w/v) SDS, (or 0.1N NaOH if for sequencing)

Neutralisation Solution:
60 mL of 5 M potassium acetate, 11.5 mL glacial acetic acid, 28.5 mL double-distilled H2O. The resulting solution is 3 M acetate and 5 M potassium and has a pH of about 4.8. Store at room temperature, and do not autoclave

Boiling Lysis Method (reference here)

This procedure is quick and reliable for preparing plasmid from small volume cultures. Even though the quality of plasmid obtained from this method is lower than that of alkaline lysis mentioned above, it is sufficient for restriction digestion.

Lysis is performed by treatment with lysozyme (0.5mg/mL final) in STET buffer and heat, which causes the chromosomal DNA to precipitate with the bacterial membrane and other proteins, which is then removed by centrifugation. The plasmid DNA is then precipitated using isopropanol.

This method is not compatible with E. coli strains that produce endonuclease enzymes, unless of course you add the additional step of a phenol:chloroform cleanup.

If you are in love with microwaves, you can also lyse the bacteria using by nuking for 20-25s in STET buffer (reference here) – don’t be too in love, things can get quite explosive in microwaves – and then simply centrifuge and precipitate with isopropanol after cooling on ice.

Reagents Required:

STET solution: 8% (w/v) sucrose, 50 mM Tris-HCl (pH 8.0), 50 mM EDTA (pH 8.0), 5
(w/v) Triton X-100. Filter-sterilize and store at 4°C.

Diatomaceous Earth Method (reference here)

For those of you keen on DIY plasmid prep kits, this one is for you

Make a suspension of 50g diatomaceous earth (Celite) in 500mL dH2O and allow to settle for three hours, then collect the solid and resuspend in at 10mg/mL in the DNA binding solution (see below).

Then working in 0.5mL volumes (scale up as needed), add 1mL DNA Binding solution, shake and leave at room temperature for 2-5 minutes to allow binding. Centrifuge for 10 seconds to pellet the particles, and resuspend the wash buffer containing. Repeat this step, then wash the pellet with 1mL of acetone and dry briefly at 65oC. Elute the DNA by adding 50-200uL and incubating at 65degC for 2 minutes.

Reagents Required:

DNA binding solution: 4M guanidine thiocyanate, 50mM Tris pH 7.0, EDTA 20mM (store in dark, stable for 3 months).
Wash buffer: 50% ethanol, 200mM NaCl, 10mM EDTA, 50mM Tris-HCl pH 7.4

Zwitterionic Lysis Method (reference here)

This is a relatively gentle procedure for lysing the cells and for plasmid isolation that relies on the soft aggregation of cellular proteins which is known to contain a large amount of plasmid DNA (which is supposedly lost in other methods). Contaminants and RNA are simply removed using wash steps, and the plasmid is eluted with TE or water.

Centrifuged cells are re-suspended in Tris (50mM), EDTA(10mM) buffer pH 8.0 and allowed to incubate for at least 1 minute in an equal volume of 0.2M NaOH and 10µM 3-(dodecyldimethyl-ammonio) propane sulfonate. Prior to centrifugation, 1mL of wash buffer comprising 10mM Tris (pH 8.0) and 10mM NaCl is added to the aggregate, after which the sample is centrifuged at max speed, most of the supernatant is removed carefully (takes some getting used to), and the wash step is either repeated, or the plasmid is eluted with TE or dH2O and the protein aggregate then discarded.

Liam’s Plasmid Method

Ok, so this hasn’t been published, and no I haven’t tested it, but I’ll bet a pair of old socks that it works like a bomb. I thought whilst writing this piece, that a combination of these methods that is quick, cheap and not too dirty would be helpful. I have also added a tweak using arginine in the lysis buffer that has been reported to give better quality DNA. So this is what I’ve come up with:

1, Spin down your pellet and resuspend pellet in 1/3 the original culture volume with STE buffer, vortex to resuspend and wash, then spin down again at maximum speed.

2. Resuspend by vortexing your pellet in 3/10 the original culture volume with buffer 1.

3. Add an equal volume of L-arginine lysis buffer. This can be left for longer than the normal 5 minutes with no problem (but not obviously overnight), mix well, but do not vortex. This high, but controlled pH solution should result in less denatured plasmid and prevent co-precipitation with protein and peptidoglycan fragments, resulting in better restriction digests later.

4. Neutralise the solution with an equal volume of 5M potassium acetate (pH 4.75) (Lithium and Sodium acetate work as well, provided they are made with equimolar quantities of acetate salt and acetic acid)

5. Spin the lysate down at maximum speed for 10 minutes then transfer the supernatant new tube and the precipitate the plasma DNA with 0.8 volumes of isopropanol which can then either be chilled at -20degC for 15 minutes, or spun straight away at max speed

Reagents Required:

Buffer 1: 50mM Tris, 10mM EDTA pH 8.0, 100ug/mL RNase A

L-arginine lysis buffer: 1% SDS solution containing 0.5M L-arginine (pH 11.7)

Neutralization solution: 5M potassium acetate (pH 4.75)

The L-arginine lysis buffer itself can be used as a substitute for the lysis buffer in commercial column-based methods (remember to recycle the columns!) and will give an increased yield and a better quality DNA

Although there is a small cost addition with the use of the L-arginine buffer, the amount of time and expense saved by eliminating failure of enzyme reactions and sequencing makes it worth it.

Please let me know if this method works for you, and yes, I will be trying it in the next few weeks and will post comparative results right here – so watch this space!

12 Comments

  1. Kyle on April 20, 2010 at 4:02 am

    I’ve tried the Arginine buffer for about a month now and just thought I’d report that I’m consistently getting lower yields and lower 260/230 ratios (usually < 1.5)on a nanodrop. I've pHed the 0.5 M Arg around 12, so unless there's something I'm missing, I'm going back to NaOH. I always like trying new things though (it makes me feel like a scientist).

  2. Liam on February 26, 2010 at 6:54 pm

    Yes, a filter could be good. Perhaps something like a funnel/tube with a sintered glass bottom that could be cleaned with HCl once the plasmid prep has been done. I suppose one would just need to tinker to figure something out that works. As far as I remember, the detergent wasn’t very expensive, and you don’t use very much.

  3. Andy on February 26, 2010 at 5:18 pm

    Maybe if they combined it with some cheesecloth to filter through or something it might work better. I might try it eventually, but that’d mean ordering some of the detergent, which I’m reluctant to do.

  4. Liam on February 24, 2010 at 6:55 am

    Hi Andy

    I didn’t go into more detail as I found the method a little difficult to work with. The protein aggregate is not solid and stuck to the tube, but rather a floating mass that you’re supposed to elute your DNA from. Perhaps with practice it might be a good method, but I found it a little to “soft” to work with. For the washing, I would guess 2-3 volumes of the protein aggregate, but like I said, only a guess.

    Good luck if you try it

  5. Andy on February 24, 2010 at 12:02 am

    That zwitterionic detergent method sounds damn interesting, but the paper is near incomprehensible – probably why you didn’t go into more detail on it. If I can get transfection-grade DNA with a simple, cheap protocol, I’ll jump on it.

    For example, they talk about adding wash buffer to the aggregate, but don’t say how much of it should be added to how much lysate, or whether it’s just made to 10mm Tris/10mM NaCl final in the lysate. Nor do they summarize what conditions they determined to be optimal. Anyone used this successfully and got a protocol?

  6. Kyle on May 13, 2009 at 10:04 pm

    Here’s another that I recently found:

    A nonalkaline method for isolating sequencing-ready plasmids
    Analytical Biochemistry, Volume 377, Issue 2, 15 June 2008, Pages 218-222
    Bonnie Paul, Cheri Cloninger, Marilyn Felton, Ronik Khachatoorian, Stan Metzenberg

    A little easier then alkaline lysis. Just pellet, add lysis buffer (3M lithium acetate (pH4.8), 0.5% SDS, 2 mM EDTA), resuspend, take a 5 minute break, spin 10 minutes, alcohol precipitate the supernatant.

    My biggest problem is that the lithium acetate won’t stay in solution ):

  7. Michael Redd on January 29, 2009 at 10:09 pm

    To Liam.
    I am interested in cheap clean DNA miniprep methods. Did you ever compare the methods in your article?
    To MolBioMonk
    Can you send your PEG-8000 precipitation protocol?

  8. Liam on July 29, 2008 at 1:38 pm

    I’ll do a comparison of precipitation methods on a later post with some actual results, as there seems to be no consensus as to which is the best for plasmid preps, and I’ll look into a high throughput adaptation of the above.

  9. jonathan on July 29, 2008 at 1:01 pm

    Nice! IMHO this is exactly the sort of science blogging we need more of – practical, well written guides for bench protocols. Looking forward to see more of this sort of thing.

  10. MolBioMonk on July 29, 2008 at 11:29 am

    Nice overview, our sequencing lab mostly uses the alkaline lysis method, and i personally precipitate plasmid in PEG-8000 buffer instead of isopropanol. We usually grow our E. coli in 96 deepwell plates and try to do the plasmid isolation in 96 well plates for high throughput. Most annoying step there is to get your supernatant out after spinning down the lysate. Any thoughts on optimizing minipreps for high-throughput?

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