Best plasmid miniprep?
Posted 06 September 2009 - 09:42 AM
I've been having trouble digesting my plasmid - it won't digest. I've used the classic "easyprep" protocol and another alkaline lysis method involving PEG. Both are undigestable and I've been trying to see some results for the past two months now with no luck. I'm thinking about trying another protocol, any ideas?
Posted 06 September 2009 - 04:55 PM
Posted 06 September 2009 - 05:01 PM
Failing that, I have had great success with the classic alkaline lysis protocol followed by isopropanol precipitation, with a later EtOH (re)precipitation and 3 x 70% EtOH washes. But try using LB if you aren't already.
Hi phage, how's it going?
Edited by swanny, 06 September 2009 - 05:01 PM.
Posted 06 September 2009 - 06:15 PM
Almost certainly the problem with rich medium is overload of the columns or lysis buffer capacity. If you just used less volume in the miniprep with the rich medium, the problem would go away.
Posted 06 September 2009 - 07:45 PM
1.Innoculate 5 ml of YT + Amp
2. Grow for about 20 hours (overnight-ish)
3. Pellet cells
4. Resuspend in resuspension buffer (containing tris, EDTA, glucose)
5. Incubate 10 minutes at room temperature
6. Add lysis solution (containing SDS and NaOH)
7. Incubate for 10 minutes at room temperature
8. Add NaOAc (pH 5.2)
9. Incubate 15 minutes on ice
10. Spin at full speed for 5 minutes
11. Transfer supernatant to new tubes
12. Add PEG solution
13. Incubate on ice for 30 minutes
14. Spin at full speed for 15 minutes
15. Remove supernatant
16. Wash with 95% EtOH (300 ul)
17. Spin down for 5 minutes
18. Remove EtOH by pipetting
19. Wash with 70% EtOH (300 ul)
20. Spin down 5 min
21. Remove EtOH, spin down and remove residual
22. Dessicate for 20 minutes
23. Resuspend in 30 ul of sterile water
I ran 3 ul on a gel and see a lot of plasmid of expected size - so the extraction works. The problem is when I digest it, I don't get anything cutting.
The plasmid I'm after is pBS and the RE I'm using is PvuII, which generates a fragment of about 800 bp without insert on pBS. Cutting pBS that I've extracted using Fermentas GeneJet generates this fragment when cut with pvuII, so it's looks like there's something in the miniprep that is inhibiting the RE. I use about 3-6 ul of sample in a 10 ul RE digestion.
I've also tried the classic "easyprep" protocol and have gotten similar results - my plasmid doesn't want to be cut, and that's when my supervisor encouraged me to use the above protocol, and it looks like I'm getting the same kind of thing.
I've looked a variety of minipreps and the PEG thing seems slightly foreign. I'm not sure what it's used for and some minipreps do virtually the same thing but leave out the PEG. WHat's with this stuff?
Posted 06 September 2009 - 08:00 PM
2 ul DNA
5 ul RE buffer
1 ul PvuII
42 ul water
digest 1 hour at 37 C
Run 20 ul on a gel along with 1 ul of the uncut DNA in an adjacent lane
Several problems arise with small volume digestions: too much DNA, too much reaction inhibitor because of the high volume of DNA to reaction volume ratio, too much glycerol from the RE storage buffer.
Posted 06 September 2009 - 08:25 PM
What prep do you typically use?
I don't see how increasing the volume up to 50 ul would help. If the problem was too much glycerol in the buffer, we're still adding the same amount (5 ul to 50 total vs. 1 ul to 10), so that shouldn't matter?
I will look more closely at the optimal RE conditions for pvuII (10 units for 1 ug for most enzymes, yes?), because maybe I'm adding too much. My supervisor gave me the go on the RE digest though, with the volumes of DNA that have lead to my failure.
But again, I haven't had trouble digesting my pBS control using the same conditions (roughly same amount of DNA, RE, etc.), it's just the miniprep stuff. And the fact that I DO see plasmid means that the prep worked, but may be dirty. Would phenoling it, and then washing it help you figure?
Posted 06 September 2009 - 09:28 PM
Posted 06 September 2009 - 10:07 PM
I'm trying to narrow it down to possible problems.
1. DNA is dirty - too high ionic strength
2. Too much DNA
3. Organic solvent carryover (EtOH - which is unlikely because my samples spent 30 minutes in the dessicator)
4. PEG. Evil PEG. I don't know what this does, but it's a goopy mess and I don't like it. It sometimes leaves an insoluble mess in my ependorff that takes a whole lot of vortexing to dissolve. Some of my ependorffs have their walls smeared with this chalky white substance - for this I blame PEG.
But back to the original topic for this thread: what's your favorite plasmid miniprep? I understand they're all pretty much the same, but some use phenol:chloroform, while others don't, some use boiling, some use PEG, some use lysozyme, SDS, etc. What do you guys have the most success with?
Posted 07 September 2009 - 05:53 AM
The main problem with your low volume digestion is that 30-60% of the volume is from your miniprep. This means that any bad things, such as ethanol, carried over proteins, PEG, lysis buffer, whatever, are present in high concentration in your restriction digest. If you do the digest I suggested, the DNA will be 4% of the volume of your reaction, and you will have a 10x or more lower concentration of those contaminants, whatever they may be. The glycerol comes not from the buffer, but from the enzyme preparation itself, so this, too, is diluted.
I would get rid of the PEG in your prep as well. Substitute potassium acetate pH 5.2 for your sodium acetate, take the supernatent and precipitate with 2.5 volumes of cold ethanol. Chill at -80 for 30 minutes, centrifuge at high speed for 20 minutes, wash with 70% ethanol. Make sure the lysis step is not overloaded with excess cells.
This will give a dirty prep, but probably good enough to work with. It sounds as if you already have clean pBS plasmid -- why do you need so much more?
Posted 07 September 2009 - 06:32 AM
Are you suggesting that I follow the miniprep procedure as I've written it, but use KOAc, leave out the PEG portion, and go directly to ethanol precipitation? Or you just suggesting a way to clean up what I already have?
Why is KOAc used instead of NaOAc?
Posted 07 September 2009 - 01:41 PM
If you are getting substantially less DNA from your prep, then you should check the antibiotic in your growth medium and figure out where the DNA is going.
The change to KOAc from NaOAc precipitates the SDS as KDS after lysis, making the isolation of genomic DNA easier. It will form a white precipitate when the lysis is neutralized.
I suggest that you don't need to clean up your existing DNA, but simply to use less of it in higher volume.
You could also clean up your DNA effectively with a phenol/chloroform extraction, but I don't recommend that as a first step without some guidance.
Posted 16 July 2011 - 09:52 AM
Posted 19 July 2011 - 11:10 PM